The Prokaryotes: Prokaryotic Physiology and Biochemistry [4 ed.] 978-3-642-30140-7, 978-3-642-30141-4

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The Prokaryotes

Eugene Rosenberg (Editor-in-Chief) Edward F. DeLong, Stephen Lory, Erko Stackebrandt and Fabiano Thompson (Eds.)

The Prokaryotes Prokaryotic Physiology and Biochemistry Fourth Edition With 220 Figures and 62 Tables

Editor-in-Chief Eugene Rosenberg Department of Molecular Microbiology and Biotechnology Tel Aviv University Tel Aviv, Israel Editors Edward F. DeLong Department of Biological Engineering Massachusetts Institute of Technology Cambridge, MA, USA Stephen Lory Department of Microbiology and Immunology Harvard Medical School Boston, MA, USA

Fabiano Thompson Laboratory of Microbiology, Institute of Biology, Center for Health Sciences Federal University of Rio de Janeiro (UFRJ) Ilha do Funda˜o, Rio de Janeiro, Brazil

Erko Stackebrandt Leibniz Institute DSMZ-German Collection of Microorganisms and Cell Cultures Braunschweig, Germany

ISBN 978-3-642-30140-7 ISBN 978-3-642-30141-4 (eBook) ISBN 978-3-642-30142-1 (print and electronic bundle) DOI 10.1007/978-3-642-30141-4 Springer Heidelberg New York Dordrecht London Library of Congress Control Number: 2012955034 3rd edition: © Springer Science+Business Media, LLC 2006 4th edition: © Springer-Verlag Berlin Heidelberg 2013 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. Exempted from this legal reservation are brief excerpts in connection with reviews or scholarly analysis or material supplied specifically for the purpose of being entered and executed on a computer system, for exclusive use by the purchaser of the work. Duplication of this publication or parts thereof is permitted only under the provisions of the Copyright Law of the Publisher’s location, in its current version, and permission for use must always be obtained from Springer. Permissions for use may be obtained through RightsLink at the Copyright Clearance Center. Violations are liable to prosecution under the respective Copyright Law. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. While the advice and information in this book are believed to be true and accurate at the date of publication, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Springer is part of Springer ScienceþBusiness Media (www.springer.com)

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 11 volumes (planned) 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 organism-oriented 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 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 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. The 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, that is, 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

Preface During most of the twentieth century, microbiologists studied pure cultures under defined laboratory conditions in order to uncover the causative agents of disease and subsequently as ideal model systems to discover the fundamental principles of genetics and biochemistry. Microbiology as a discipline onto itself, e.g., microbial ecology, diversity, and evolution-based taxonomy, has only recently been the subject of general interest, partly because of the realization that microorganisms play a key role in the environment. The development and application of powerful culture-independent molecular techniques and bioinformatics tools has made this development possible. The fourth edition of the Handbook of the Prokaryotes has been updated and expanded in order to reflect this new era of microbiology. The first five volumes of the fourth edition contain 34 updated and 43 entirely new chapters. Most of the new chapters are in the two new sections: Prokaryotic Communities and Bacteria in Human Health and Disease. A collection of microorganisms occupying the same physical habitat is called a ‘‘community,’’ and several examples of bacterial communities are presented in the Prokaryotic Communities section, organized by Edward F. DeLong. Over the last decade, important advances in molecular biology and bioinformatics have led to the development of innovative culture-independent approaches for describing microbial communities. These new strategies, based on the analysis of DNA directly extracted from environmental samples, circumvent the steps of isolation and culturing of microorganisms, which are known for their selectivity leading to a nonrepresentative view of prokaryotic diversity. Describing bacterial communities is the first step in understanding the complex, interacting microbial systems in the natural world. The section on Bacteria in Human Health and Disease, organized by Stephen Lory, contains chapters on most of the important bacterial diseases, each written by an expert in the field. In addition, there are separate general chapters on identification of pathogens by classical and non-culturing molecular techniques and virulence mechanisms, such as adhesion and bacterial toxins. In recognition of the recent important research on beneficial bacteria in human health, the section also includes chapters on gut microbiota, prebiotics, and probiotics. Together with the updated and expanded chapter on Bacterial Pharmaceutical Products, this section is a valuable resource to graduate students, teachers, and researchers interested in medical microbiology. Volumes 6–11, organized by Erko Stackebrandt and Fabiano Thompson, contain chapters on each of the ca. 300 known prokaryotic families. Each chapter presents both the historical and current taxonomy of higher taxa, mostly above the genus level; molecular analyses (e.g., DDH, MLSA, riboprinting, and MALDI-TOF); genomic and phenetic properties of the taxa covered; genome analyses including nonchromosomal genetic elements; phenotypic analyses; methods for the enrichment, isolation, and maintenance of members of the family; ecological studies; clinical relevance; and applications. As in the third edition, the volumes in the fourth edition are available both as hard copies and e-books, and as eReferences. The advantages of the online version include no restriction of color illustrations, the possibility of updating chapters continuously and, most importantly, libraries can place their subscribed copies on their servers, making it available to their community in offices and laboratories. The editors thank all the chapter authors and the editorial staff of Springer, especially Hanna Hensler-Fritton, Isabel Ullmann, Daniel Quin˜ones, Alejandra Kudo, and Audrey Wong, for making this contribution possible. Eugene Rosenberg Editor-in-Chief

About the Editors Eugene Rosenberg (Editor-in-Chief) Department of Molecular Microbiology and Biotechnology Tel Aviv University Tel Aviv Israel

Eugene Rosenberg holds a Ph.D. in biochemistry from Columbia University (1961) where he described the chemical structures of the capsules of Hemophilus influenzae, types B, E, and F. His postdoctoral research was performed in organic chemistry under the guidance of Lord Todd in Cambridge University. He was an assistant and associate professor of microbiology at the University of California at Los Angeles from 1962 to 1970, where he worked on the biochemistry of Myxococcus xanthus. Since 1970, he has been in the Department of Molecular Microbiology and Biotechnology, Tel Aviv University, as an associate professor (1970–1974), full professor (1975–2005), and professor emeritus (2006–present). He has held the Gol Chair in Applied and Environmental Microbiology since 1989. He is a member of the American Academy of Microbiology and European Academy of Microbiology. He has been awarded a Guggenheim Fellowship, a Fogarty International Scholar of the NIH, the Pan Lab Prize of the Society of Industrial Microbiology, the Proctor & Gamble Prize of the ASM, the Sakov Prize, the Landau Prize, and the Israel Prize for a ‘‘Beautiful Israel.’’ His research has focused on myxobacteriology; hydrocarbon microbiology; surface-active polymers from Acinetobacter; bioremediation; coral microbiology; and the role of symbiotic microorganisms in the adaptation, development, behavior, and evolution of animals and plants. He is the author of about 250 research papers and reviews, 9 books, and 16 patents.

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About the Editors

Edward F. DeLong Department of Biological Engineering Massachusetts Institute of Technology Cambridge, MA USA

Edward DeLong received his bachelor of science in bacteriology at the University of California, Davis, and his Ph.D. in marine biology at Scripps Institute of Oceanography at the University of California, San Diego. He was a professor at the University of California, Santa Barbara, in the Department of Ecology for 7 years, before moving to the Monterey Bay Aquarium Research Institute where he was a senior scientist and chair of the science department, also for 7 years. He now serves as a professor at the Massachusetts Institute of Technology in the Department of Biological Engineering, where he holds the Morton and Claire Goulder Family Professorship in Environmental Systems. DeLong’s scientific interests focus primarily on central questions in marine microbial genomics, biogeochemistry, ecology, and evolution. A large part of DeLong’s efforts have been devoted to the study of microbes and microbial processes in the ocean, combining laboratory and field-based approaches. Development and application of genomic, biochemical, and metabolic approaches to study and exploit microbial communities and processes is his another area of interest. DeLong is a fellow in the American Academy of Arts and Science, the U.S. National Academy of Science, and the American Association for the Advancement of Science.

About the Editors

Stephen Lory Department of Microbiology and Immunology Harvard Medical School Boston, MA USA

Stephen Lory received his Ph.D. degree in Microbiology from the University of California in Los Angeles in 1980. The topic of his doctoral thesis was the structure-activity relationships of bacterial exotoxins. He carried out his postdoctoral research on the basic mechanism of protein secretion by Gram-negative bacteria in the Bacterial Physiology Unit at Harvard Medical School. In 1984, he was appointed assistant professor in the Department of Microbiology at the University of Washington in Seattle, becoming full professor in 1995. While at the University of Washington, he developed an active research program in host-pathogen interactions including the role of bacterial adhesion to mammalian cells in virulence and regulation of gene expression by bacterial pathogens. In 2000, he returned to Harvard Medical School where he is currently a professor of microbiology and immunobiology. He is a regular reviewer of research projects on various scientific panels of governmental and private funding agencies and served for four years on the Scientific Council of Institute Pasteur in Paris. His current research interests include evolution of bacterial virulence, studies on post-translational regulation of gene expression in Pseudomonas, and the development of novel antibiotics targeting multi-drugresistant opportunistic pathogens.

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About the Editors

Erko Stackebrandt Leibniz Institute DSMZ-German Collection of Microorganisms and Cell Cultures Braunschweig Germany

Erko Stackebrandt holds a Ph.D. in microbiology from the Ludwig-Maximilians University Munich (1974). During his postdoctoral research, he worked at the German Culture Collection in Munich (1972–1977), 1978 with Carl Woese at the University of Illinois, Urbana Champaign, and from 1979 to 1983 he was a member of Karl Schleifer’s research group at the Technical University, Munich. He habilitated in 1983 and was appointed head of the Departments of Microbiology at the University of Kiel (1984–1990), at the University of Queensland, Brisbane, Australia (1990–1993), and at the Technical University Braunschweig, where he also was the director of the DSMZ-German Collection of Microorganisms and Cell Cultures GmbH (1993–2009). He is involved in systematics, and molecular phylogeny and ecology of Archaea and Bacteria for more than 40 years. He has been involved in many research projects funded by the German Science Foundation, German Ministry for Science and Technology, and the European Union, working on pure cultures and microbial communities. His projects include work in soil and peat, Mediterranean coastal waters, North Sea and Baltic Sea, Antarctic Lakes, Australian soil and artesian wells, formation of stromatolites, as well as on giant ants, holothurians, rumen of cows, and the digestive tract of koalas. He has been involved in the description and taxonomic revision of more than 650 bacteria taxa of various ranks. He received a Heisenberg stipend (1982–1983) and his work has been awarded by the Academy of Science at Go¨ttingen, Bergey’s Trust (Bergey’s Award and Bergey’s Medal), the Technical University Munich, the Australian Society for Microbiology, and the American Society for Microbiology. He held teaching positions in Kunming, China; Budapest, Hungary; and Florence, Italy. He has published more than 600 papers in refereed journals and has written more than 80 book chapters. He is the editor of two Springer journals and served as an associate editor of several international journals and books as well as on national and international scientific and review panels of the German Research Council, European Science Foundation, European Space Agency, and the Organisation for Economic Co-Operation and Development.

About the Editors

Fabiano Thompson Laboratory of Microbiology Institute of Biology Center for Health Sciences Federal University of Rio de Janeiro (UFRJ) Ilha do Funda˜o Rio de Janeiro Brazil

Fabiano Thompson became the director of research at the Institute of Biology, Federal University of Rio de Janeiro (UFRJ), in 2012. He was an oceanographer at the Federal University of Rio Grande (Brazil) in 1997. He received his Ph.D. in biochemistry from Ghent University (Belgium) in 2003, with emphasis on marine microbial taxonomy and biodiversity. Thompson was an associate researcher in the BCCM/LMG Bacteria Collection (Ghent University) in 2004; professor of genetics in 2006 at the Institute of Biology, UFRJ; and professor of marine biology in 2011 at the same university. He has been a representative of UFRJ in the National Institute of Metrology (INMETRO) since 2009. Thompson is the president of the subcommittee on the Systematics of Vibrionaceae–IUMS and an associate editor of BMC Genomics and Microbial Ecology. The Thompson Lab in Rio currently performs research on marine microbiology in the Blue Amazon, the realm in the southwestern Atlantic that encompasses a variety of systems, including deep sea, Cabo Frio upwelling area, Amazonia river-plume continuum, mesophotic reefs, Abrolhos coral reef bank, and Oceanic Islands (Fernando de Noronha, Saint Peter and Saint Paul, and Trindade).

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Table of Contents Physiology and Biochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .1 1 Acetogenic Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Harold L. Drake . Kirsten Ku¨sel . Carola Matthies

2 Virulence Strategies of Plant Pathogenic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 Maeli Melotto . Barbara N. Kunkel

3 Oxidation of Inorganic Nitrogen Compounds as an Energy Source . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Eberhard Bock . Michael Wagner

4 H2-Metabolizing Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 Edward Schwartz . Johannes Fritsch . Ba¨rbel Friedrich

5 Hydrocarbon-Oxidizing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 Eugene Rosenberg

6 Lignocellulose-Decomposing Bacteria and Their Enzyme Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 Edward A. Bayer . Yuval Shoham . Raphael Lamed

7 Aerobic Methylotrophic Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 Ludmila Chistoserdova . Mary E. Lidstrom

8 Dissimilatory Fe(III)- and Mn(IV)-Reducing Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 Derek Lovley

9 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 309 Ralf Rabus . Theo A. Hansen . Friedrich Widdel

10 Denitrifying Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405 James P. Shapleigh

11 Dinitrogen-Fixing Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 Ernesto Ormen˜o-Orrillo . Mariangela Hungria . Esperanza Martinez-Romero

12 Magnetotactic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453 Dennis A. Bazylinski . Christopher T. Lefe`vre . Dirk Schu¨ler

13 Luminous Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 495 Paul V. Dunlap . Henryk Urbanczyk

14 Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 529 Dimitry Y. Sorokin . Horia Banciu . Lesley A. Robertson . J. Gijs Kuenen . M. S. Muntyan . Gerard Muyzer

15 Colorless Sulfur Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 555 Gerard Muyzer . J. Gijs Kuenen . Lesley A. Robertson

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Table of Contents

16 Bacterial Stress Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 589 Eliora Z. Ron

17 Anaerobic Biodegradation of Hydrocarbons Including Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 605 Johann Heider . Karola Schu¨hle

18 Physiology and Biochemistry of the Methane-Producing Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 635 Reiner Hedderich . William B. Whitman

List of Contributors Horia Banciu Faculty of Biology and Geology Babes-Bolyai University Cluj-Napoca Romania

Johannes Fritsch Institut fu¨r Biologie, Mikrobiologie, Humboldt-Universita¨t zu Berlin Berlin Germany

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

Theo A. Hansen Microbial Physiology (MICFYS) University of Groningen Groningen The Netherlands

Dennis A. Bazylinski School of Life Sciences University of Nevada at Las Vegas Las Vegas, NV USA

Reiner Hedderich Max Planck Institute fu¨r Terrestriche Mikrobiologie Marburg Germany

Eberhard Bock Institute of General Botany Department of Microbiology University of Hamburg Hamburg Germany

Johann Heider Fachbereich Biologie Laboratorium fu¨r Mikrobiologie Marburg Germany

Ludmila Chistoserdova Department of Chemical Engineering University of Washington Seattle, WA USA Harold L. Drake Department of Ecological Microbiology BITOEK University of Bayreuth Bayreuth Germany Paul V. Dunlap University of Michigan Ann Arbor, MI USA Ba¨rbel Friedrich Institut fu¨r Biologie, Mikrobiologie, Humboldt-Universita¨t zu Berlin Berlin Germany

Mariangela Hungria Embrapa Soja Londrina Brazil J. Gijs Kuenen Department of Biotechnology Delft University of Technology Delft The Netherlands Barbara N. Kunkel Department of Biology Washington University St. Louis, MO USA Kirsten Ku¨sel Friedrich Schiller University Jena Institute of Ecology Limnology/Aquatic Geomicrobiology Jena Germany

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List of Contributors

Raphael Lamed Department of Molecular Microbiology and Biotechnology George S. Wise Faculty of Life Sciences Tel Aviv University Ramat Aviv Israel Christopher T. Lefe`vre CEA Cadarache/CNRS/Universite´ Aix-Marseille II, UMR7265 Service de Biologie Ve´ge´tale et de Microbiologie Environnementale Laboratoire de Bioe´nerge´tique Cellulaire Saint Paul lez Durance France Mary E. Lidstrom Department of Chemical Engineering and Department of Microbiology University of Washington Seattle, WA USA Derek Lovley Department of Microbiology University of Massachusetts Amherst, MA USA Esperanza Martinez-Romero Genomic Sciences Center, UNAM Cuernavaca Mexico Carola Matthies Department of Ecological Microbiology BITOEK University of Bayreuth Bayreuth Germany Maeli Melotto Department of Biology University of Texas Arlington, TX USA M. S. Muntyan Belozersky Institute of Physico-Chemical Biology Moscow State University Moscow Russia

Gerard Muyzer Department of Biotechnology Delft University of Technology Delft The Netherlands and Department of Aquatic Microbiology Institute for Biodiversity and Ecosystem Dynamics University of Amsterdam Amsterdam The Netherlands Ernesto Ormen˜o-Orrillo Genomic Sciences Center, UNAM Cuernavaca Mexico Ralf Rabus Institute for Chemistry and Biology of the Marine Environment (ICBM) University of Oldenburg Oldenburg Germany Lesley A. Robertson Department of Biotechnology Delft University of Technology Delft The Netherlands Eliora Z. Ron Department of Molecular Microbiology and Biotechnology The George S. Wise Faculty of Life Sciences Tel Aviv University Tel Aviv Israel Eugene Rosenberg Department of Molecular Microbiology and Biotechnology Tel Aviv University Tel Aviv Israel Karola Schu¨hle Fachbereich Biologie Laboratorium fu¨r Mikrobiologie Marburg Germany Dirk Schu¨ler Department Biologie I Ludwig-Maximilians-Universita¨t Mu¨nchen Planegg-Martinsried Germany

List of Contributors

Edward Schwartz Institut fu¨r Biologie, Mikrobiologie, Humboldt-Universita¨t zu Berlin Berlin Germany

Henryk Urbanczyk University of Miyazaki Miyazaki City, Miyazaki Japan

James P. Shapleigh Department of Microbiology Cornell University Ithaca, NY USA

Michael Wagner Department of Microbial Ecology Faculty Center of Ecology Faculty of Life Sciences University of Vienna Vienna Austria

Yuval Shoham Department of Biotechnology and Food Engineering Technion – Israel Institute of Technology Haifa Israel Dimitry Y. Sorokin Winogradsky Institute of Microbiology Russian Academy of Sciences Moscow Russia and Department of Biotechnology Delft University of Technology Delft The Netherlands

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

Friedrich Widdel Max-Planck-Institut fu¨r Marine Mikrobiologie Bremen Germany

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Physiology and Biochemistry

1 Acetogenic Prokaryotes Harold L. Drake1 . Kirsten Ku¨sel 2 . Carola Matthies1 1 Department of Ecological Microbiology, BITOEK, University of Bayreuth, Bayreuth, Germany 2 Friedrich Schiller University Jena, Institute of Ecology, Limnology/Aquatic Geomicrobiology, Jena, Germany

Introduction to Acetogenic Bacteria and the Process of Acetogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acetogens Defined . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Usage of the Terms ‘‘Acetogenesis,’’ ‘‘Homoacetogen,’’ and ‘‘Homoacetogenesis’’ . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Global Impact and Evolutionary Perspectives . . . . . . . . . . . .

3 4 4 5

Historical Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Discovery of Acetogenic Bacteria and Acetogenesis . . . . . . . 6 Resolution of the Acetyl-CoA ‘‘Wood/Ljungdahl’’ Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Isolates to Date and Microbiological Methods . . . . . . . . . . . . . . 8 Description of Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Cultivation Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 Taxonomy and Phylogeny . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 Detection of Acetogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 The Acetyl-CoA Pathway and Bioenergetics . . . . . . . . . . . . . . . . 26 CO2 as Terminal Electron Acceptor and the Concept of Fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 Enzymology of the Acetyl-CoA Pathway . . . . . . . . . . . . . . . . . 29 Conservation of Energy and Bioenergetics . . . . . . . . . . . . . . . 31 Occurrence of the Acetyl-CoA Pathway in Nonacetogenic Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Diverse Physiological Talents of Acetogens . . . . . . . . . . . . . . . . . 33 Diverse Electron Donors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 Use of Diverse Terminal Electron Acceptors . . . . . . . . . . . . . 36 Regulation of the Acetyl-CoA Pathway and Other Metabolic Abilities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 37 Tolerance to Oxic Conditions and Metabolism of O2 . . . 38 Ecology of Acetogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39 Metabolic Interactions of Acetogens in Pure Cultures and Complex Ecosystems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 39 Diverse Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41 Biotechnological Applications of Acetogens . . . . . . . . . . . . . . . . 47 Commercial Production of Acetic Acid from Sugars . . . . 47 Bioconversion of Synthesis Gas to Acetic Acid, Ethanol, and Other Chemicals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48 Bioremediation, Bioreactors, and Landfills . . . . . . . . . . . . . . 48 Other Potential Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48 Summary and Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48

Abstract This chapter circumscribes the acetogens, a physiologically defined group of the domain Bacteria that are anaerobes, using the acetyl-CoA pathway as a mechanism for the reductive synthesis of acetyl-CoA from CO2, for a terminal-electronaccepting, energy-conserving process, and for mechanism for the fixation (assimilation) of CO2 in the synthesis of cell carbon. Three main metabolic features of these organisms were defined, such as the use of chemolithoautotrophic substrates (H2-CO2 or CO-CO2) as sole sources of carbon and energy under anoxic conditions, the capacity to convert certain sugars stoichiometrically to acetate, and the ability to O-demethylate methoxylated aromatic compounds and metabolize the O-methyl group via the 420 acetylCoA pathway. Acetogens have been assigned to more than 20 different genera and they differ in their morphology, cytology, and physiology. The most frequently isolated acetogenic species to date are members of the genera Clostridium and Acetobacterium. The habitat, the morphological and physiological properties, and the phylogenetic position of acetogenic species are presented. The electron flow of the “Wood/Ljungdahl” pathway as well as properties and function of enzymes involved in the acetylCoA pathway is shown in detail. Several biotechnological applications are described with the commercial production of acetic acid from sugars and the bioconversion of synthesis gas to acetic acid, ethanol, and other chemicals being the most important ones.

Introduction to Acetogenic Bacteria and the Process of Acetogenesis This chapter presents an overview of the history, taxonomy, phylogenetics, biochemistry, physiology, ecology, and applied aspects of acetogens. Acetogenic prokaryotes have only been found in the domain bacteria. These prokaryotes utilize a reductive onecarbon pathway for the synthesis of acetyl-CoA, a metabolic precursor of both acetate and biomass. This pathway fixes CO2 and is termed ‘‘the acetyl-CoA pathway.’’ This pathway is often referred to as ‘‘the Wood/Ljungdahl pathway’’ in recognition of the two individuals, Harland G. Wood and Lars G. Ljungdahl, who were responsible for elucidating most of its enzymological features from the model acetogen Moorella thermoacetica (> Fig. 1.1; see the section on > ‘‘Historical Perspectives’’ in this chapter). Acetogenesis (i.e., the process by which acetogens synthesize acetate) is often regarded as a fermentation process; however, as outlined in the section on > ‘‘CO2 as Terminal Electron Acceptor and the Concept of Fermentation,’’ acetogenesis is

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_61, # Springer-Verlag Berlin Heidelberg 2013

4

1

Acetogenic Prokaryotes

a

b

Harland G. Wood

c

0.5 µm

Lars G. Ljungdahl

. Fig. 1.1 (a) Electron micrograph of a sporulated cell of Clostridium thermoaceticum, which was reclassified as Moorella thermoacetica (Collins et al. 1994). From Drake (1994), used with permission from Kluwer Academic. (b, c) The two biochemists who were primarily responsible for resolving the enzymological features of the acetyl-CoA ‘‘Wood/Ljungdahl’’ pathway in M. thermoacetica. From Drake and Daniel (2004), used with permission from Elsevier. The dates of the photos for B and C are September 1977 (taken after Harland Wood’s 70th birthday celebration/symposium at Case Western Reserve University, during which Wood was ‘‘roasted’’ and given the honorary degree of Doctor of Mouse Science [the image on the hood symbolizes the shape of transcarboxylase as seen by electron microscopy]) and May 2000 (taken during the symposium honoring Harry D. Peck, Jr., at the University of Georgia), respectively

very dissimilar to classic fermentations. Purinolytic bacteria that synthesize acetate via the glycine pathway will not be considered in this chapter. However, certain features of this CO2-fixing, glycine-reductase-dependent pathway are similar to those of the acetyl-CoA pathway, and the reader is directed to the review of Andreesen (1994) for a detailed assessment of this pathway and organisms that use it.

Acetogens Defined Usage of the term ‘‘acetogen’’ has not been consistent in the literature, and this inconsistent usage has caused a small amount

of confusion regarding which organisms utilize the acetyl-CoA pathway for the synthesis of acetate. The following definition for the term acetogen has been previously proposed (Drake 1994) and is applied in this chapter: Acetogen: An anaerobe that can use the acetyl-CoA pathway as a (1) mechanism for the reductive synthesis of acetyl-CoA from CO2, (2) terminal-electron-accepting, energy-conserving process, and (3) mechanism for the fixation (assimilation) of CO2 in the synthesis of cell carbon. Per this definition, the formation of acetate as an end product is unimportant, i.e., the fate of acetyl-CoA is less important than the process by which it is formed. For example, Eubacterium limosum, ‘‘Butyribacterium methylotrophicum,’’ and Caloramator pfennigii (formerly Clostridium pfennigii), organisms that qualify as acetogens per the above definition, form butyrate from the acetyl-CoA that is formed via the acetyl-CoA pathway (Lynd and Zeikus 1983; Zeikus 1983; Krumholz and Bryant 1985; Zeikus et al. 1985; Loubiere et al. 1992). Likewise, the acetogen Acetobacterium woodii forms ethanol from acetyl-CoA under certain conditions (Buschhorn et al. 1989). The term ‘‘acetogenic’’ is an adjective that could be used to describe any organism that makes acetate or acetic acid. However, the metabolic processes by which acetate can be formed during either the aerobic or anaerobic growth of diverse microorganisms might not be equivalent. The mechanism by which acetate is formed via the oxidation of ethanol by Acetobacter aceti is fundamentally different from that used by certain obligate anaerobes that synthesize acetate from CO2 via the reductive acetyl-CoA pathway. Thus, it is important that a differential nomenclature be applied to distinguish between acetateforming bacteria because failure to do so results in unnecessary confusion in the literature. For example, Thermobacteroides proteolyticus and the syntroph PA-1 have been referred to as acetogens because they form acetate from glucose (Ollivier et al. 1985b; Brulla and Bryant 1989). However, these organisms use protons, not CO2, as terminal electron acceptors and form H2, not acetate, as their main reduced end product; in short, they do not appear to use the acetyl-CoA pathway for the synthesis of acetate. Likewise, the butyrate-degrading syntroph Syntrophomonas wolfei has been described as an acetogen (Stams and Dong 1995). However, this organism (1) converts butyrate to acetate and H2 (which can subsequently be used to reduce CO2 to formate) by -oxidation via the crotonyl-CoA pathway (Wofford et al. 1986), (2) does not reduce CO2 to acetate, and (3) is not known to utilized the acetyl-CoA pathway.

Usage of the Terms ‘‘Acetogenesis,’’ ‘‘Homoacetogen,’’ and ‘‘Homoacetogenesis’’ The term ‘‘acetogenesis’’ could be used to describe the process by which any organism forms acetate. For example, the term acetogenesis has been used to describe the (1) oxygen-dependent process by which Enterococcus RfL6 oxidizes lactate to acetate (Tholen et al. 1997) and (2) the production of acetate during

Acetogenic Prokaryotes

proteolysis by Treponema denticola (Mikx 1997). No evidence suggests that these organisms utilize the acetyl-CoA pathway for acetate synthesis. Since such usage makes it difficult to understand what process is being referred to, it has been suggested that usage of the term acetogenesis be restricted to processes by which two molecules of CO2 are used to form one molecule of acetate (Wood and Ljungdahl 1991). Unfortunately, such usage fails to adequately distinguish between the three known metabolic processes by which acetate is formed from CO2: (1) the acetyl-CoA pathway, (2) the glycine-synthase-dependent pathway, and (3) the reductive citric acid cycle (Fuchs 1986, 1989; Thauer 1988; Wood and Ljungdahl 1991). The term ‘‘homoacetogen’’ is often used to distinguish between organisms that use the acetyl-CoA pathway and those that do not (Schink and Bomar 1992). This term implies that acetate is the sole product formed by a particular organism. However, organisms that use the acetyl-CoA pathway usually do not form acetate as their sole end product. Their capacity to form any particular end product, including acetate, is dependent upon cultivation conditions. Butyrate (Lynd and Zeikus 1983; Krumholz and Bryant 1985; Worden et al. 1989; Grethlein et al. 1991), ethanol (Buschhorn et al. 1989), lactate (Lorowitz and Bryant 1984; Drake 1993; Misoph and Drake 1996a), succinate (Dorn et al. 1978; Lorowitz and Bryant 1984; Matthies et al. 1993; Misoph and Drake 1996a), reduced aromatic acrylates (Tschech and Pfennig 1984; Parekh et al. 1992; Misoph et al. 1996b), reduced aromatic aldehydes (Lux et al. 1990), CO (Diekert et al. 1986), H2 (Martin et al. 1983; Lorowitz and Bryant 1984; Savage et al. 1987), CH4 (Savage et al. 1987; Buschhorn et al. 1989), sulfide (Heijthuijsen and Hansen 1989; Beaty and Ljungdahl 1991), dimethylsulfide (Beaty and Ljungdahl 1991), nitrite (Seifritz et al. 1993; Fro¨stl et al. 1996), and ammonium (Seifritz et al. 1993, 2003; Fro¨stl et al. 1996) are examples of reduced end products of so-called homoacetogens. Indeed, the production of such products can constitute the sole energy-conserving, growth-supportive process of the cell (see the section on > ‘‘Use of Diverse Terminal Electron Acceptors’’ in this chapter). Thus, the conditions under which an acetogen forms acetate should be qualified rather than merely referring to the organism as a homoacetogen. For example, Ruminococcus productus (formerly Peptostreptococcus productus) is homoacetogenic on pyruvate but forms acetate, lactate, succinate, and formate when cultivated on fructose; this acetogen can also form large amounts of ethanol during glycerol-dependent growth (Misoph and Drake 1996a). Likewise, Moorella thermoacetica is homoacetogenic when cultivated on H2/CO2 but does not form acetate when cultivated on H2/CO2 in the presence of nitrate; under this condition, the dissimilation of nitrate is used preferentially to acetogenesis for the conservation of energy (Fro¨stl et al. 1996). Lastly, the term ‘‘homoacetate production’’ has been used to describe the process by which a genetically modified strain of Escherichia coli anaerobically produces 2 moles of acetate per mole glucose fermented (Causey et al. 2003), yet this process is not homoacetogenic (i.e., does not yield 3 moles of acetate per mole glucose) and the acetyl-CoA pathway is not involved.

1

Independent of these problems of usage, the production of acetate as the sole end product from certain sugars, H2/CO2, or CO/CO2 strongly suggests that the organism in question utilizes the acetyl-CoA pathway per the definition for the term acetogen (see the section on > ‘‘Acetogens Defined’’ in this chapter).

Global Impact and Evolutionary Perspectives Acetogens were initially viewed as obscure, poorly defined microorganisms. For nearly five decades following the discovery of acetogens in the 1930s, the major interest in them was restricted to resolving the biochemical features of the acetyl-CoA pathway (see the section on > ‘‘Historical Perspectives’’ in this chapter). The microbiology of acetogens drew little interest until the 1980s when it started to become apparent that acetogens were a widely distributed, phylogenetically diverse group of microorganisms. Added interest in the acetyl-CoA pathway occurred when it was discovered that methanogens and sulfate-reducing bacteria used metabolic pathways that contained acetyl-CoA synthase, one of the key enzymes in the acetyl-CoA pathway (Fuchs 1986, 1989; Schauder et al. 1986; Thauer et al. 1989; see the section on > ‘‘Occurrence of the Acetyl-CoA Pathway in Nonacetogenic Microorganisms’’ in this chapter). Major bacterial groups employing this pathway in either the direction of acetate/biomass synthesis or acetate degradation include acetogens, methanogens, and sulfate-reducing bacteria. It is not possible to determine how much carbon is processed globally via acetogens and pathways that make use of acetyl-CoA synthase. However, several facts are noteworthy: 1. The Calvin cycle, the reductive tricarboxylic acid cycle, the hydroxypropionate cycle, and the acetyl-CoAWood/Ljungdahl pathway facilitate the complete autotrophic fixation of CO2. Of these pathways, the one-carbon acetyl-CoA pathway is biochemically the most simple. For example, the acetyl-CoA pathway requires less ATP to fix a molecule CO2 than does the Calvin cycle. Furthermore, the acetyl-CoA pathway is a linear process that does not depend on preformed, complex molecules to which CO2 is fixed in a cyclic process (e.g., the Calvin cycle, the reductive tricarboxylic acid cycle, the hydroxypropionate cycle are dependent upon ribulose biphosphate, oxalacetate, and acetyl-CoA, respectively, for the fixation of CO2) (see section on > ‘‘The Acetyl-CoA Pathway and Bioenergetics’’ in this chapter). Methanogens utilize an acetyl-CoA-synthase-dependent pathway that is biochemically very similar to the acetyl-CoA pathway utilized by acetogenic bacteria (see the section on > ‘‘Occurrence of the Acetyl-CoA Pathway in Nonacetogenic Microorganisms’’ in this chapter), and methanogens (or ancestors of methanogens) may have been the first autotrophs (Schopf et al. 1983; Brock 1989). Thus, and since life originated under anoxic conditions, the acetyl-CoA pathway or a pathway closely related to it may have been the first process used for the autotrophic fixation of CO2 (Fuchs 1986, Wood and Ljungdahl 1991; Lindahl and Chang 2001).

5

6

1

Acetogenic Prokaryotes

2. Approximately half of the human population contains low numbers of methanogens in their gastrointestinal systems and produces relatively little CH4; the colon of these individuals, as well as of those who more actively emit CH4, is heavily colonized by acetogens (Wolin and Miller 1983). Indeed, the gastrointestinal systems of mammals, whether they harbor methanogens or not, are heavily colonized with acetogens (Prins and Lankhorst 1977; Breznak and Kane 1990; Mackie and Bryant 1994; Wolin and Miller 1994; Leedle et al. 1995). 3. Acetogens inhabit the human colon. In this habitat, acetogens produce 1010 kg of acetate per year from H2-CO2, and acetogenesis is one of the dominant processes in the overall metabolism of carbohydrate in the human colon (Lajoie et al. 1988; Wolin and Miller 1994; Dore´ et al. 1995; Bernalier et al. 1996a, b; Miller and Wolin 1996; Wolin et al. 1999). 4. Totally, 1012 kg of acetate are produced each year via the reduction of CO2 by acetogens in the hindgut of termites, a number that is fivefold greater than the annual amount of methane formed globally via the biogenic reduction of CO2 (Breznak and Kane 1990). One-third of the energy requirements of the termite is provided by the acetate that is synthesized by the reduction of CO2 by gut acetogens (Breznak 1994). 5. Totally 1013 kg of acetate is formed and further metabolized annually in terrestrial habitats such as soils and sediments, and a minimum of 10 % of this acetate is likely formed by the reduction of CO2 via the acetyl-CoA pathway (Wood and Ljungdahl 1991). 6. Up to 25 % of the total organic carbon of soil can be turned over through acetate under low temperature, anoxic conditions (equivalent to nearly 40 g acetate per kg dry wt. of soil; Ku¨sel and Drake 1994). The capacity to form acetate in soils is concomitant with acetogenic activities and the occurrence of H2-utilizing acetogens (Ku¨sel and Drake 1995; Wagner et al. 1996; Ku¨sel et al. 1999c). Acetate is a dominant organic compound in soil solution (Tani et al. 1993), and concentrations can be in the mM range following a rainfall event (Ku¨sel and Drake 1999a). Assuming a weight of 1017 kg for the first meter of the global terrestrial surface [based on a surface area of 1014 m2 (Whitman et al. 1998) and using a weight conversion of 103 kg per m3] and an acetate concentration of 0.1 mmol per kg of this material, it can be estimated that 1012 kg of acetate is present in the first meter of the terrestrial surface at any one moment (i.e., per ‘‘snapshot’’). Even if only a small percentage of this acetate were formed by acetogens, given the turnover dynamics of acetate, the annual magnitude of the acetogen-derived acetate in the terrestrial biosphere would be enormous. The number of prokaryotes in the terrestrial subsurface might exceed that of the terrestrial surface by a factor of 10 (Whitman et al. 1998). It can be projected that acetate and acetogens are also involved in the cycling of carbon in this poorly explored compartment of the terrestrial ecosphere (see the section on > ‘‘Diverse Habitats’’ in this chapter).

7. The acetate formed by acetogenesis is an essential trophic link during the turnover of carbon in diverse anoxic habitats (McInerney and Bryant 1981). Such observations not only illustrate that nature’s ability to form acetate is enormous, they also demonstrate that the acetylCoA Wood/Ljungdahl pathway is fundamental to the carbon cycle of earth.

Historical Perspectives Discovery of Acetogenic Bacteria and Acetogenesis Acetogenesis was first reported in 1932, when unknown organisms in sewage were shown to catalyze the H2-dependent reduction of CO2 to acetate (Fischer et al. 1932). Shortly thereafter, the Dutch microbiologist Wieringa reported the isolation of the first acetogen (Wieringa 1936, 1939–1940, 1941). The organism, Clostridium aceticum, was a sporeforming, mesophilic rod and grew at the expense of the following reaction: 4H2 þ 2CO2 ! CH2 COOH þ 2H2 O

ð1:1Þ

Such a reaction had not been observed earlier. With the exception of a small study on the nutritional requirements of C. aceticum (Karlsson et al. 1948), no further work was published with this acetogen until it was reisolated in 1980–1981 (Adamse 1980; Braun et al. 1981; Gottschalk and Braun 1981; > Fig. 1.2). Clostridium thermoaceticum was discovered a few years after the isolation of C. aceticum (Fontaine et al. 1942) and was the only acetogen available for laboratory study for several decades (> Fig. 1.1). This bacterium was reclassified as Moorella thermoacetica (Collins et al. 1994) and will be referred to by this name hereafter. Moorella thermoacetica was isolated as an obligate heterotroph and was observed to convert glucose to acetate; the stoichiometry of this process approximated the following reaction: C6 H12 O6 ! 3CH3 COOH

ð1:2Þ

In the early 1940s, no known metabolic process could explain this reaction, and it was proposed that the CO2 produced via oxidation was subsequently utilized in the synthesis of acetate: Since, in this fermentation, 2.5 moles of a two-carbon compound (acetic acid) are obtained from 1 mole of glucose, it seems probable that either there is some primary cleavage of glucose other than the classical 3-3 split or that a one-carbon compound is being reabsorbed. Of these two possibilities, the recent work on carbon dioxide uptake makes the latter seem more likely (Fontaine et al. 1942). The latter statement was in reference to the discovery of CO2 fixation in heterotrophs (Wood and Werkman 1936, 1938; Wood et al. 1941a, b). Subsequent proposals for the acetogenic conversion of glucose or pyruvate to acetate made it possible to

Acetogenic Prokaryotes

a

1

The overlap between reactions > 1.1, > 1.4, and > 1.7 indicated that a unique reductive process was likely responsible for acetate synthesis from CO2.

b

Resolution of the Acetyl-CoA ‘‘Wood/Ljungdahl’’ Pathway Barker and Kamen (1945) demonstrated in the first published biological experiments with 14C (Kamen 1963) that M. thermoacetica incorporated 14CO2 equally into both carbon atoms of acetate. This landmark experiment with 14C demonstrated that the capacity of M. thermoacetica to synthesize acetate from glucose was, in fact, similar to the capacity of C. aceticum to synthesize acetate from H2/CO2: "

1µm

. Fig. 1.2 (a). Tube containing dried soil and spores of the first acetogen to be isolated, Clostridium aceticum. The tube was obtained from H. A. Barker; the date on the tube is May 7, 1947. B. Electron micrograph of a peritrichously flagellated cell of C. aceticum [From Braun et al. (1981), used with permission from Springer. The photograph (panel A) was kindly provided by G. Gottschalk]

see that both the autotrophic and heterotrophic acetogenic processes likely involved the reductive synthesis of acetate from CO2 (Barker 1944). Conversion of glucose to acetate: Oxidative portion: C6 H12 O6 þ 2H2 O ! 2CH3 COOH þ 2CO2 þ 8H

ð1:3Þ Reductive portion: 8H þ 2CO2 ! CH3 COOH þ 2H2 O ð1:4Þ Net reaction: C6 H12 O6 ! 3CH3 COOH

ð1:5Þ

Conversion of pyruvate to acetate: Oxidative portion: 4C3 H4 O3 þ 4H2 O ! 4CH3 COOH þ 4CO2 þ 8H

ð1:6Þ Reductive portion: 8H þ 2CO2 ! CH3 COOH þ 2H2 O ð1:7Þ Net reaction: 4C3 H4 O3 þ 2H2 O ! 5CH3 COOH þ 2CO2 ð1:8Þ

It may be concluded that the acetic acid fermentation of glucose by C. thermoaceticum involves a partial oxidation of the substrate to 2 moles each of acetic acid and carbon dioxide followed by a reduction and condensation of the carbon dioxide to a third mole of acetic acid. (Barker and Kamen 1945)

In 1952, Wood repeated the 14C experiments of Barker and Kamen with 13CO2 and confirmed that M. thermoacetica synthesized acetate from two molecules of CO2 (Wood 1952a). In this work, mass spectrometry conclusively demonstrated that CO2 was uniformly fixed into both the carboxyl and methyl carbons of the third molecule of acetate from glucose. Utilizing [3,4-14C]-glucose, it was also shown that carbons 3 and 4 of glucose were converted to CO2 (Wood 1952b). These early studies by Barker, Kamen, and Wood demonstrated that (1) glucose was subject to a classic 3-3 split between carbons 3 and 4 and (2) CO2 was fixed via an unknown CO2-fixing process into acetate (> Fig. 1.3). It took decades of continued research before the enzymology of this CO2-fixing process was fully resolved (see the section on > ‘‘The Acetyl-CoA Pathway and Bioenergetics’’ in this chapter). It is an irony of the history of acetogenesis that the model organism (i.e., M. thermoacetica) used to resolve the biochemistry of this autotrophic process was thought to be an obligate heterotroph during these decades of research. Indeed, the chemolithoautotrophic nature of M. thermoacetica (Daniel et al. 1990) was resolved nearly five decades after its isolation and well after the enzymological details of the acetyl-CoA pathway were firmly established. The milestones of the numerous studies that resolved both the enzymology of the acetyl-CoA pathway and the chemolithoautotrophic abilities of the model acetogen used in these studies can be found in numerous review articles (Ljungdahl and Wood 1969; Wood 1972, 1976, 1982, 1985, 1989, 1991; Ljungdahl 1986; Wood and Ljungdahl 1991; Drake 1992, 1994; Ragsdale 1991, 1994, 1997) and are outlined in > Table 1.1. For additional insights into the early career years of Harland G. Wood, see the recent excellent historical treatments by Singleton (Singleton 1997a, b, 2000).

7

8

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a

. Table 1.1 Milestones that led to resolving the acetyl-CoA pathway and chemolithoautotrophic abilities of Moorella thermoacetica

Glucose Glycolysis

2 ATPSLP 4 [H]

b 2 Pyruvate Pruvateferredoxin Oxidoreductase

2 CO2 4 [H]

2 Acetyl-CoA

2 CO2

Acetate Kinase 2 Acetate

Eventa

1932

H2-dependent conversion of CO2 to acetate in sewage sludge (Fischer et al. 1932)

1936

Isolation of the first acetogen, Clostridium aceticum; total synthesis of acetate from H2-CO2 (Note: culture was lost, Wieringa 1936, 1939–1940)

1942

Discovery of the second acetogen, Moorella thermoacetica (formerly Clostridium thermoaceticum); conversion of one glucose to three acetate molecules (Fontaine et al. 1942)

1944

Acetogenic conversion of pyruvate to acetate (Barker 1944)

c

Phosphotransacetylase 2 Acetylphosphate

Year

Acetyl-CoA Pathway

8 [H]

2 ATPSLP

1945–1952 Synthesis of acetate from 14CO2 (Barker and Kamen 1945) or 13CO2 (Wood 1952a, b) 1955

Formate as a methyl-group precursor (Lentz and Wood 1955)

1964

Methylcobalamin as methyl-group precursor (Poston et al. 1964)

1965

Autotrophic synthesis of cell-carbon precursors from CO2 (Ljungdahl and Wood 1965)

Acetate

. Fig. 1.3 Homoacetogenic conversion of glucose to acetate. Glucose is first converted to two molecules of pyruvate via glycolysis (Box A); glycolysis yields ATP by substrate-level phosphorylation (SLP). Pyruvate is then oxidized and decarboxylated, yielding acetyl-CoA, CO2, and reducing equivalents (Box B). The two acetylCoA molecules that are produced from pyruvate are converted to two molecules of acetate; this process yields additional ATP by SLP. The eight reducing equivalents that are produced via glycolysis and pyruvate-ferredoxin oxidoreductase are utilized in the acetyl-CoA pathway to reduce two molecules of CO2 to an additional molecule of acetate (Box C). The CO2 that is reduced in the acetyl-CoA pathway is likely derived primarily from supplemental CO2 rather than the CO2 derived via the decarboxylation of pyruvate (Modified from Drake 1994)

1966–1969 Proposal of one-carbon pathway for the tetrahydrofolate/corrinoid-mediated synthesis of acetate from CO2 (Ljungdahl and Wood 1966, 1969) 1973–1986 Resolution of the tetrahydrofolate pathway [reviewed in Ljungdahl (1986)] 1978–1980 Discovery of CO dehydrogenase as a nickelcontaining enzyme (Diekert and Thauer 1978; Drake et al. 1980) 1981

Resolution of enzymes required for synthesis of acetyl-CoA from pyruvate and methyltetrahydrofolate (Drake et al. 1981a)

1981–1982 Demonstration that CO replaces the carboxyl group of pyruvate and undergoes an exchange reaction with acetyl-CoA (Drake et al. 1981b; Hu et al. 1982) 1982

Discovery of hydrogenase (Drake 1982)

Isolates to Date and Microbiological Methods

1983

Purification of CO dehydrogenase (Diekert and Ritter 1983; Ragsdale et al. 1983)

The number of known acetogens has increased significantly in the last two decades, and approximately 100 different species have been isolated to date from extremely diverse habitats. Acetogens can be found in almost all anoxic environments, including some extreme habitats, as indicated by the isolation of strain SS1 (Liu and Suflita 1993) and ‘‘Acetobacterium psammolithicum’’ from deep subsurface sediment and sandstone, respectively (Krumholz et al. 1999). Although most isolates to date are mesophilic, thermophilic and psychrotolerant species have also been isolated. The occurrence and ecological roles of acetogens in various habitats are discussed in the section on acetogen ecology (see the section on > ‘‘Ecology of Acetogens’’ in this chapter).

1983

Use of H2 and CO under organotrophic conditions (Kerby and Zeikus 1983)

1984

Resolution of nutritional requirements (Lundie and Drake 1984)

1984

Enzyme system for H2-dependent synthesis of acetyl-CoA (Pezacka and Wood 1984b)

1984–1986 CO dehydrogenase is acetyl-CoA synthase (Pezacka and Wood 1984a, b; Regsdale and Wood 1985), and CO is the carbonyl precursor in the acetyl-CoA pathway under growth conditions (Diekert et al. 1984; Martin et al. 1985) 1985–1991 Catalytic mechanism of acetyl-CoA synthase [reviewed in Ragsdale (1991)]

Acetogenic Prokaryotes

. Table 1.1 (continued) Year

Eventa

1986–1990 H2- and CO-dependent electron transport system coupled to the synthesis of ATP (Ivey and Ljungdahl 1986; Hugenholtz and Ljungdahl 1989, 1990; Das et al. 1989) 1990

Chemolithoautotrophic growth on H2-CO2 and CO-CO2 (Daniel et al. 1990)

1991

Integrated model for catabolic, anabolic, and bioenergetic features of the acetyl-CoA ‘‘Wood/ Ljungdahl’’ pathway (Wood and Ljungdahl 1991)

a Events prior to the isolation of Moorella thermoacetica Modified from Drake (1994)

Bacteria considered to be acetogens as defined above (see the section on > ‘‘Acetogens Defined’’ in this chapter) are listed in > Table 1.2. However, relatively few of these bacteria have been examined in detail, and a good understanding of the metabolic capabilities of most of the isolates is lacking. In compiling the list, the apparent acetogenic capability (see the section on > ‘‘Usage of the Terms ‘Acetogenesis,’ ‘Homoacetogen,’ and ‘Homoacetogenesis’’’ in this chapter) of each organism has been taken into account. In this regard, three main metabolic features of these organisms (Drake 1994) are (1) the use of chemolithoautotrophic substrates (H2-CO2 or CO-CO2) as sole sources of carbon and energy under anoxic conditions, (2) the capacity to convert certain sugars stoichiometrically to acetate, and (3) the ability to O-demethylate methoxylated aromatic compounds and metabolize the O-methyl group via the acetyl-CoA pathway. Many acetogens display all three of these metabolic capabilities. Most acetogenic isolates are rod-shaped, but coccoid forms have also been observed (> Table 1.2). Staining properties vary, sometimes within a genus, and both Gram-negative and Gram-positive species have been reported (> Table 1.2). Some acetogens have flagella and are motile. Some form spores that remain viable for long periods; the thermophilic sporeformers are fairly resistant to high temperatures. Indeed, spores of M. thermoacetica have a decimal reduction time (i.e., the time required to decrease the population of viable spores by 90 %) of 111 min at 121  C (Byrer et al. 2000). Cells of the acetogen Clostridium glycolicum RD-1 are tethered by connecting filaments, a morphological structure recently described for Clostridium akagii and Clostridium uliginosum (Kuhner et al. 2000; Matthies et al. 2001). Thus, the ultrastructural features of acetogens are highly variable.

Description of Species Acetogens have been assigned to 21 different genera and differ in their morphological, cytological, and physiological properties (> Table 1.2). The genera Clostridium and Acetobacterium

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harbor the most acetogenic species isolated to date. The first acetogen was classified as a clostridial species, C. aceticum (Wieringa 1936). The second acetogenic genus Acetobacterium was established when the first Gram-positive, nonsporeforming acetogen (Acetobacterium woodii; Balch et al. 1977; > Fig. 1.4) was isolated and could not be grouped with the acetogenic clostridia. Conspicuously, all the heretofore isolated psychrotolerant acetogens and many N2-fixing acetogens belong to the genus Acetobacterium (Schink and Bomar 1992; > Table 1.2). About half of the genera that harbor acetogens only contain one acetogenic species (e.g., Holophaga foetida, Acetohalobium arabaticum, Oxobacter pfennigii, Acetonema longum; > Table 1.2). Recently, acetogenesis has been observed in spirochetes (‘‘Treponema primitia’’) isolated from termite guts (Leadbetter et al. 1999; Graber and Breznak 2004a; Graber et al. 2004b; > Fig. 1.5). A brief overview of acetogenic species having validated names is given in the following paragraphs. The names of those organisms not validated are in quotation marks. Earlier compilations include Breznak (1992), Diekert (1992), Hippe et al. (1992), Schink and Bomar (1992), Mackie and Bryant (1994), and Schink (1994). Although relatively few of the acetogens listed below have been evaluated for their ability to tolerate O2, it should be anticipated that many acetogens possess the ability to both tolerate and consume small amounts of oxygen (Ku¨sel et al. 2001; Karnholz et al. 2002; Boga and Brune 2003). Acetitomaculum ruminis. This species was isolated from steer rumen fluid (Greening and Leedle 1989). Cells are Gram-positive, nonsporeforming, motile, slightly curved rods. Growthsupportive substrates include H2-CO2, CO, formate, cellobiose, glucose, ferulate, and syringate. With all substrates, acetate is the sole reduced end product (Greening and Leedle 1989). Acetoanaerobium noterae. This species was isolated from sediment samples of the Notera oil exploration site in Israel (Sleat et al. 1985). Cells are Gram-positive, nonsporeforming, motile, straight rods. Acetoanaerobium noterae grows with H2-CO2, glucose, and maltose and produces acetate as the sole product. Propionate, butyrate, isobutyrate, and isovalerate are also formed when yeast extract serves as the growthsupportive substrate (Sleat et al. 1985). Acetoanaerobium romashkovii. This organism was isolated from the Romashkino oil field in Tatarstan (DavydovaCharakhch’yan et al. 1992). Cells are Gram-positive, nonsporeforming, motile rods with rounded ends. Growth-supportive substrates include H2-CO2, formate, methanol, pyruvate, lactate, ethylene glycol, sugars, and amino acids. Acetate is the sole product from carbohydrates and H2-CO2; propionate is also formed during growth on sucrose. ‘‘Acetoanaerobium romashkovii’’ produces and excretes polysaccharides during growth on H2-CO2 or methanol (Davydova-Charakhch’yan et al. 1992). Acetobacterium bakii, Acetobacterium fimetarium, and Acetobacterium paludosum. These species were isolated from cold habitats ( Fig. 1.4). Growth-supportive substrates include H2-CO2, CO, formate, methanol, 2,3-butandiol, ethylene glycol, acetoin, glycerol, sugars, betaine, and several methoxylated aromatic acids (Balch et al. 1977; Bache and Pfennig 1981; Eichler and Schink 1984; Sharak Genthner and Bryant 1987; Schink and Bomar 1992). Cultures demethylate the osmolytes dimethylsulfoniopropionate and glycine-betaine to methylthiopropionate and dimethylglycine, respectively; however, only the demethylation of glycine-betaine supported growth of the organism (Jansen and Hansen 2001). Acetobacterium woodii growths mixotrophically on (i.e., can simultaneously utilize) H2-CO2 and organic compounds (e.g., fructose; Braun and Gottschalk 1981) and can use aromatic acrylates as energy-conserving, growth-supportive terminal electron acceptors (Bache and Pfennig 1981; Tschech and Pfennig 1984). Growth, motility, and acetate formation from H2-CO2 are strictly dependent on sodium ions (Heise et al. 1989; Mu¨ller and Bowien 1995; Aufurth et al. 1998). Several Na+-dependent reactions in the metabolism of A. woodii have been identified, and associated enzymes have been purified and characterized (Heise et al. 1989, 1991, 1992, 1993; Mu¨ller and Gottschalk 1994; Reidlinger and Mu¨ller 1994a; Reidlinger et al. 1994b; Mu¨ller et al. 2001). Cells reductively dechlorinate carbon tetrachloride (Egli et al. 1988; Stromeyer et al. 1992); dechlorination is enhanced by the addition of hydroxocobalamin (Hashsham and Freedman 1999). Cells also tolerate and consume small amounts of oxygen (Karnholz et al. 2002). Acetohalobium arabaticum. This organism was isolated from a cyanobacterial mat in a saline lagoon and was the first obligately halophilic acetogen to be described (Zhilina and Zavarzin 1990). Sodium chloride (10–25 %) is necessary for growth. Cells are motile, straight rods often aggregated in palisades. H2-CO2, CO, trimethylamine, formate, betaine, lactate, pyruvate, and histidine are growth-supportive substrates. Acetate is the main product during growth on trimethylamine and betaine and is accompanied by minor amounts of methylamines (Zhilina and Zavarzin 1990; Zavarzin et al. 1994). Cell extracts have CO dehydrogenase and hydrogenase activities, which are stimulated by increased salt concentrations.

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Acetonema longum. This organism was isolated from the gut contents of the wood-feeding termite Pterotermes occidentis (Kane and Breznak 1991a). Cells are sporeforming, motile rods of unusually large size; cells can be up to 60 mm in length. Growth-supportive substrates include H2-CO2, pyruvate, fumarate, glucose, mannitol, and ribose; poor growth occurs on citrate, propanol, ethylene glycol, and 3,4,5-trimethoxybenzoate. Homoacetogenesis only occurs with H2-CO2. Butyrate and acetate are the main products from carbohydrates and pyruvate; fumarate is metabolized to propionate and acetate, and rhamnose yields 1,2-propanediol as the major product (Kane and Breznak 1991a). Bryantella formatexigens. This species was isolated from human feces (Wolin et al. 2003). Cells are Gram-positive, nonmotile short rods (approx. 1.2  0.7 mm). Single cells, and pairs and short chains of cells, are apparent. Upon isolation, the type strain (I-52; Wolin and Miller 1994) fermented vegetable cellulose and carboxymethylcellulose but lost this ability after storage under frozen conditions. No growth occurs on H2-CO2 or formate, and formate is required for optimal homoacetogenic conversion of glucose. The lack of supplemental formate yields succinate, lactate, and acetate as products from glucose. These characteristics indicate that the formate dehydrogenase is negligible. Growth is supported by stachyose, sucrose, lactose, maltose, galactose, mannose, and xylose. Cells are catalase and oxidase negative, and nitrate is not reduced. Butyribacterium methylotrophicum. This organism was isolated from a sewage digestor (Zeikus et al. 1980). Cells are Grampositive, sporeforming, nonmotile rods. Growth is supported by H2-CO2, formate, methanol, glucose, fructose, sucrose, pyruvate, lactate, and glycerol (Zeikus et al. 1980; Kerby and Zeikus 1987). Homoacetogenic utilization of substrates only occurs with H2-CO2 and formate. With other substrates, butyrate and H2 are also produced (Zeikus et al. 1980; Lynd and Zeikus 1983). After prolonged incubation in medium with CO in the gas phase, the type strain grew on and utilized CO; acetate was the sole product from CO (Lynd et al. 1982). There is substantial evidence that ‘‘B. methylotrophicum’’ and Eubacterium limosum are the same species: (1) the metabolic properties of the two organisms are nearly identical and (2) the 16S rRNA gene sequences of the two organisms are very similar (99.4% sequence similarity; Moore and Cato 1965; Sharak Genthner et al. 1981; Tanner et al. 1981; Sharak Genthner and Bryant 1982; Tanner and Woese 1994; Jansen and Hansen 2001). Caloramator fervidus. This species was isolated from a hot spring in New Zealand and was first described as Clostridium fervidus (Patel et al. 1987). Cells are Gram-negative, sporeforming, motile rods. Carbohydrates support growth, and acetate is the major end product. However, growth on one-carbon compounds (e.g., formate) or other typical acetogenic substrates (e.g., H2-CO2) has not been reported, and substrate/product stoichiometries of carbohydrate utilization are not available. Thus, the true acetogenic nature of this organism has not been established. Until otherwise proven, one should assume that the organism might not be an acetogen.

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Acetogenic Prokaryotes

Clostridium aceticum. This species was the first acetogen to be isolated (> Fig. 1.2). It was isolated from soil and described by Wieringa (Wieringa 1936, 1939–1940). After early studies with the organism (Karlsson et al. 1948), it was lost for about 30 years. However, C. aceticum was reisolated from soil using Wieringa’s enrichment procedure, and almost at the same time, spores of the original Wieringa strain in sterile dried soil were found in Barker’s laboratory and revived (Adamse 1980; Braun et al. 1981). Cells are Gram-negative, sporeforming, motile rods (Wieringa 1939–1940; Braun et al. 1981). Growth-supportive substrates include H2-CO2, CO, fructose, glutamate fumarate, pyruvate, aldehyde groups of aromatic compounds, and methoxylated aromatic compounds (Wieringa 1939–1940; Braun et al. 1981; Lux and Drake 1992; Matthies et al. 1993; Go¨ßner et al. 1994). As with C. formicoaceticum (see below), fumarate is dismutated by C. aceticum to acetate and succinate and is metabolized independent of the acetyl-CoA pathway; fumarate also serves as an alternative electron acceptor and is reduced to succinate (Matthies et al. 1993). N2 is fixed (Cato et al. 1986). Clostridium autoethanogenum. This organism was isolated from rabbit feces (Abrini et al. 1994). Cells are Gram-positive, sporeforming, motile rods. The range of substrates includes H2-CO2, CO, pyruvate, hexoses, pentoses, and glutamate and is similar to the range of substrates used by Clostridium ljungdahlii (Abrini et al. 1994; Tanner et al. 1993). CO is converted to acetate and ethanol (Abrini et al. 1994). Ethanol production from CO was also reported for C. ljungdahlii; however, this metabolic potential is not necessarily stable (Barik et al. 1988; Tanner et al. 1993). The 16S rRNA gene sequences of ‘‘C. autoethanogenum’’ and C. ljungdahlii are essentially identical (Stackebrandt et al. 1999). Clostridium coccoides. Two acetogenic strains of C. coccoides (strains 1410 and 3110) were isolated from the human intestinal tract (Kamlage et al. 1997). The type strain of C. coccoides isolated from mouse feces was not initially described as an acetogen; however, it has recently been shown to contain all the enzymes of the acetyl-CoA pathway when grown on H2-CO2-formate (Kaneuchi et al. 1976; Kamlage et al. 1997). Cells of C. coccoides strain 1410 (which is probably identical to strain 3110) are Gram-variable, coccoid rods. Clostridium coccoides strain 1410 grows on a variety of hexoses, pentoses, sugar alcohols, H2-CO2-formate, and H2-CO2-vanillate. Products from growth have not been reported. However, resting cells convert formate, H2-CO2, and O-methyl groups of vanillate to acetate at stoichiometries indicative of acetogenesis; the aromatic ring of vanillate remains intact (Kamlage et al. 1997). Resting cells of C. coccoides strain 1410 convert glucose to acetate, succinate, and D-lactate. Clostridium difficile. Five acetogenic strains of C. difficile were isolated from the rumen of newborn lambs; strain AA1 is considered as a representative strain (Rieu-Lesme et al. 1998 in this chapter). No acetogenic potentials have been documented for the type strain of C. difficile. Cells of C. difficile strain AA1 are Gram-positive, sporeforming, giant filamentous rods. Growth of strain AA1 is supported by H2-CO2, fructose, glucose,

cellobiose, maltose, mannose, and syringate. Acetate is the sole product from H2-CO2, and the substrate/product stoichiometry is indicative of acetogenesis; however, glucose and fructose are metabolized to almost equal amounts of acetate and butyrate, and small amounts of ethanol and isovalerate (Rieu-Lesme et al. 1998). Clostridium formicoaceticum. The first strain of C. formicoaceticum was probably isolated from pond sediment by El Ghazzawi (1967). Although the organism was called Clostridium aceticum in the title of the German publication, El Ghazzawi stated that his isolate differed from C. aceticum and tentatively named his organism ‘‘Clostridium formicoaceticum’’ because it produced both formate and acetate (El Ghazzawi 1967). The type strain of C. formicoaceticum was isolated from sewage sludge (Andreesen et al. 1970). Cells are Gram-negative, sporeforming, motile, straight or slightly curved rods. The range of substrates is very similar to that of C. aceticum (see above) but also includes glycerol, gluconate, glucuronate, and glycerate (Andreesen et al. 1970). Clostridium formicoaceticum can be differentiated from C. aceticum by its inability to grow with H2-CO2 and its ability to grow with methanol and lactate (Andreesen et al. 1970; Lux and Drake 1992). As with C. aceticum, the utilization of fumarate by C. formicoaceticum does not involve the acetyl-CoA pathway; fumarate is dismutated to acetate and succinate (Dorn et al. 1978). Fumarate can also serve as an alternative electron acceptor (Matthies et al. 1993), and N2 is fixed (Bogdahn et al. 1983). Reductant derived from the oxidation of the aldehyde groups of certain aromatic compounds (e.g., 4-hydroxybenzaldehyde) is growth supportive (Go¨ßner et al. 1994), preferentially used in the acetylCoA pathway, and inhibits the use of fructose (Frank et al. 1998). Clostridium glycolicum. Two acetogenic strains of C. glycolicum have been isolated. Strain 22 was isolated from sewage sludge, grows on H2-CO2, and produces mainly acetate; cells are Gram-positive rods that form oval, subterminal spores (Ohwaki and Hungate 1977). Strain 22 has been deposited at the American Type Culture Collection (ATCC) and has been identified as a strain of Clostridium glycolicum; however, the 16S rRNA gene sequence is not available (per information from the ATCC Bacteriology Program). Strain RD-1 was isolated from sea grass roots and was identified as an acetogenic strain of C. glycolicum by analysis of the 16S rRNA gene sequence (Ku¨sel et al. 2001). Cells of strain RD-1 are Gram-positive, sporeforming, motile rods that can be linked by connecting filaments. Growth-supportive substrates of strain RD-1 include H2-CO2, formate, pyruvate, lactate, ethylene glycol, and certain sugars. Except for growth on sugars and ethylene glycol, acetate is the sole reduced end product. Strain RD-1 is aerotolerant and grows at O2 concentrations of up to 6 % in the headspace of static liquid cultures and up to 4 % in the headspace of shaken liquid cultures; ethanol, lactate, and H2 are the reduced end products under oxic conditions (Ku¨sel et al. 2001; see the section on > ‘‘Tolerance to Oxic Conditions and Metabolism of O2’’). No acetogenic potentials have been found for the type strain of C. glycolicum (Gaston and Stadtman 1963; Ku¨sel et al. 2001).

Acetogenic Prokaryotes

. Fig. 1.6 Electron micrograph of cells from a young culture (16 h, fructose grown) of Clostridium ljungdahlii (ATCC 55383T) with peritrichously inserted flagella. Bar equals 1 mm (The micrograph was kindly provided by R.S. Tanner)

Clostridium ljungdahlii. This organism was isolated from chicken manure/waste (Barik et al. 1988; Tanner et al. 1993). Cells are Gram-positive, sporeforming, motile rods (> Fig. 1.6). The organism grows autotrophically on H2-CO2 and CO; heterotrophic growth occurs on formate, ethanol, pyruvate, fumarate, and sugars (including fructose and xylose; Tanner et al. 1993). The sole product from H2-CO2 and fructose is acetate; however, from synthesis gas (a mixture of H2, CO, and CO2), acetate and ethanol are produced (Tanner et al. 1993; Phillips et al. 1994). Nitrate is reduced to ammonium; however, unlike the dissimilation of nitrate by M. thermoacetica (see the sections on > ‘‘Use of Diverse Terminal Electron Acceptors’’ and > ‘‘Regulation of the Acetyl-CoA Pathway and Other Metabolic Abilities’’ in this chapter), the reduction of nitrate does not have a regulatory effect on acetogenesis and likewise does not enhance the growth of the organism (Seifritz et al. 1993; Fro¨stl et al. 1996; Laopaiboon and Tanner 1999). Clostridium magnum. This species was isolated from pasteurized freshwater sediment (Schink 1984). Cells are Grampositive, sporeforming, motile, large straight rods. H2-CO2, formate, methanol, 2,3-butandiol, acetoin, malate, citrate, and a few sugars are substrates, and acetate is the sole reduced end product. N2 is fixed (Bomar et al. 1991), and small amounts of O2 are tolerated and consumed (Karnholz et al. 2002). Clostridium mayombei. This organism was isolated from the gut of a soil-feeding termite (Kane et al. 1991b). Cells are Grampositive, sporeforming, motile, straight rods. Growth occurs on H2-CO2, sugars, sugar alcohols, organic acids, and amino acids. The main reduced end product is acetate; however, succinate is metabolized to CO2 and propionate (Kane et al. 1991b). Clostridium methoxybenzovorans. This species was isolated from an olive mill wastewater digester (Mechichi et al. 1999). Cells of C. methoxybenzovorans are Gram-positive, sporeforming, nonmotile rods. Growth occurs on H2-CO2,

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methanol, lactate, sugars, methoxylated aromatic compounds, betaine, dimethylglycine, dimethylsulfide, casaminoacids, and peptone. H2-CO2 is metabolized to acetate and formate. Metabolism of betaine, dimethylglycine, and dimethylsulfide yields acetate, and sugars are metabolized to acetate, formate, ethanol, H2, and CO2. O-methyl groups, methanol, and lactate are metabolized to acetate and butyrate (Mechichi et al. 1999). Since no substrate/product stoichiometries have been reported for the organism, the acetogenic utilization of most substrates is uncertain. Clostridium scatologenes. An acetogenic strain of C. scatologenes (SL1) was isolated from sediment of an acidic coal mine pond (Ku¨sel et al. 2000). The type strain of C. scatologenes was isolated from soil and was not originally described as an acetogen (Holdeman et al. 1977). However, both strain SL1 and the type strain utilize H2 and CO with the concomitant production of acetate, and cell extracts of both organisms have CO dehydrogenase, hydrogenase, and formate dehydrogenase activities (Ku¨sel et al. 2000). Cells are Grampositive, sporeforming, motile, long rods and produce skatole, a dung odor component (Holdeman et al. 1977; Ku¨sel et al. 2000). Substrates include fructose, arabinose, ethanol, formate, vanillate, H2-CO2, and CO. The major reduced end product is acetate. However, in addition to acetate, butyrate and traces of H2 are also produced from sugars (Ku¨sel et al. 2000). Clostridium ultunense. This species was isolated from an anaerobic acetate-oxidizing triculture that was enriched from a digester fed with swine manure (Schnu¨rer et al. 1994, 1996). Cells are Gram-positive, sporeforming rods that change cell size, cell form, and motility during growth. The only known growthsupportive substrates are formate, betaine, glucose, pyruvate, ethylene glycol, and cysteine. The main end products are acetate, formate, and traces of H2. H2-CO2 does not support growth; however, H2-CO2 is converted to acetate by resting cells (Schnu¨rer et al. 1996). Acetate is oxidized in coculture with a methanogen, and the oxidation of acetate appears to occur via a reversal of the acetyl-CoA pathway (Schnu¨rer et al. 1997). An acetogen (strain AOR) that also oxidized acetate in coculture with a methanogen was previously isolated (Lee and Zinder 1988); however, this strain has been lost (S.H. Zinder, personal communication). Eubacterium aggregans. This organism was isolated from an olive mill wastewater digestor (Mechichi et al. 1998). Cells are Gram-positive, nonsporeforming, nonmotile rods that form aggregates. Substrates include H2-CO2, glucose, fructose, sucrose, lactate, formate, methanol, betaine, and numerous methoxylated aromatic compounds. Although E. aggregans is described as homoacetogenic, H2, formate, acetate, and butyrate are produced from sugars (Mechichi et al. 1998). Acetate is the sole reduced end product with formate and methanol. Methoxylated aromatic compounds are O-demethylated, and acetate, butyrate, and the corresponding hydroxylated aromatic compounds are formed. Aldehyde groups of methoxylated aromatic compounds are oxidized to carboxylate groups. Eubacterium limosum. This species was isolated from sheep rumen and digester sludge (Sharak Genthner et al. 1981). Cells

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Acetogenic Prokaryotes

are Gram-positive, nonsporeforming, nonmotile straight rods that become more pleomorphic after prolonged incubation. Eubacterium limosum is metabolically very versatile; its substrate range includes sugars, amino acids, methoxylated aromatic compounds, glycine, betaine, lactate, methanol, H2-CO2, and CO (Sharak Genthner et al. 1981; Sharak Genthner and Bryant 1982, 1987; Jansen and Hansen 2001). Both acetate and butyrate are produced from one-carbon compounds (Sharak Genthner et al. 1981; Pacaud et al. 1985; see ‘‘Butyribacterium methylotrophicum,’’ above). Cultures demethylate the osmolytes dimethylsulfoniopropionate and glycine-betaine to methylthiopropionate and dimethylglycine, respectively; however, only the demethylation of glycine-betaine supports growth of the organism (Jansen and Hansen 2001). Holophaga foetida. This organism (strain TMBS4) was isolated from freshwater sediment (Bak et al. 1992; Liesack et al. 1994). Cells are Gram-negative, nonsporeforming, nonmotile rods. The substrate range is rather small and mainly consists of pyruvate and aromatic compounds, especially methylated and nonmethylated trihydroxybenzenes. Acetate is the main reduced end product. In contrast to other acetogens, H. foetida degrades aromatic rings to acetate (Bak et al. 1992; Kreft and Schink 1993). Dimethylsulfide and methanediol are produced from methoxylated aromatic compounds when cells are cultured in sulfide-containing media, indicating that sulfide can serve as a methyl acceptor (Bak et al. 1992). CO2 and CO can also be used as methyl acceptors with the subsequent formation of acetate. CO dehydrogenase activity is present in cells grown on methoxylated aromatic compounds (Kreft and Schink 1993). Holophaga foetida occupies a fairly isolated position in the phylogenetic tree of the bacteria (Liesack et al. 1994, 1997; see the section on > ‘‘Taxonomy and Phylogeny’’). Moorella glycerini. This species is a thermophilic acetogen and was isolated from the sediment of a hot spring at Yellowstone National Park (Slobodkin et al. 1997). The cells are Gram-positive, sporeforming, motile, straight rods. Growth is supported by glycerol, sugars, lactate, glycerate, pyruvate, and yeast extract; however, H2-CO2 is not growth supportive. Acetate is the only product from glycerol and glucose. Fumarate is reduced to succinate, and the reduction of thiosulfate yields elemental sulfur. Nitrate is not dissimilated. Optimum growth occurs at 58  C. Moorella mulderi. This organism is a thermophilic acetogen and was isolated from a high-temperature bioreactor (Balk et al. 2003). The cells are Gram-positive, sporeforming rods. Growth is supported by H2-CO2, formate, methanol, hexoses, cellobiose, lactate, and pyruvate. The reduction of thiosulfate yields sulfide. Nitrate is not dissimilated. Moorella thermoacetica. This organism is a thermophilic acetogen that was isolated from horse manure and was first described as Clostridium thermoaceticum (Fontaine et al. 1942). On the basis of phylogenetic analysis of the 16S rRNA gene sequence, C. thermoaceticum was reclassified as M. thermoacetica (Collins et al. 1994). Although the organism was originally isolated from horse manure, the organism is a common inhabitant of soils (Go¨ßner and Drake 1997; Go¨ßner et al. 1998, 1999;

Karita et al. 2003). Cells are Gram-variable, sporeforming, variably motile, straight rods (> Fig. 1.1). The optimum temperature of growth is 55–60  C (Fontaine et al. 1942), and the vitamin nicotinic acid is required for growth (Lundie and Drake 1984). Moorella thermoacetica was the first bacterium that was shown to produce 3 moles of acetate from 1 mole of hexose (Fontaine et al. 1942) and is one of the most metabolically robust acetogens characterized to date. Moorella thermoacetica was originally isolated as an obligate heterotroph (Fontaine et al. 1942), but nearly five decades later, it was shown to be capable of autotrophic growth (Daniel et al. 1990). This bacterium displays very diverse physiological capabilities (Drake and Daniel 2004). Growth-supportive substrates include CO, H2-CO2, formate, methanol, hexoses, pentoses, methoxylated benzoic acids, and several two-carbon compounds (e.g., oxalate, glycolate, and glyoxylate; Fontaine et al. 1942; Daniel et al. 1990, 2004; Daniel and Drake 1993; Drake et al. 1997; Seifritz et al. 1999; Kim et al. 2002). Carboxyl groups of aromatic compounds can serve as CO2 equivalents in the acetyl-CoA pathway (Hsu et al. 1990a, b). Thiosulfate (Beaty and Ljungdahl 1990, 1991), nitrate (Seifritz et al. 1993), and nitrite (Seifritz et al. 2003) serve as alternative electron acceptors. Nitrate is dissimilated to both nitrite and ammonium, and nitrite is dissimilated to ammonium. Ethanol and n-propanol are oxidized and are growth-supportive substrates when nitrate is dissimilated; neither ethanol nor n-propanol is utilized as an acetogenic substrate (Fro¨stl et al. 1996). Reductively dechlorinates carbon tetrachloride (Egli et al. 1988). Tolerates and consumes small amounts of oxygen (Karnholz et al. 2002). A recent isolate that is phylogenetically nearly identical to M. thermoacetica is cellulolytic (Karita et al. 2003). Moorella thermoacetica is the most studied acetogen, and the enzymology of the acetyl-CoA pathway was resolved with this organism (see the section on > ‘‘Historical Perspectives’’ and > Table 1.1 in this chapter). Moorella thermoautotrophica. This organism is a thermophilic acetogen that was isolated from a hot spring at Yellowstone National Park and was first described as Clostridium thermoautotrophicum (Wiegel et al. 1981). On the basis of phylogenetic analysis of the 16S rRNA gene sequence, C. thermoautotrophicum was reclassified as M. thermoautotrophica (Collins et al. 1994). Cells are Gram-variable, sporeforming, motile rods (Wiegel et al. 1981). Moorella thermoautotrophica was initially described as being metabolically distinct from the closely related M. thermoacetica (Collins et al. 1994); this distinction was primarily based on the H2-dependent acetogenic abilities of the former bacterium (Wiegel et al. 1981). However, later studies demonstrated that M. thermoacetica grows chemolithoautotrophically on H2-CO2 (Daniel et al. 1990). Both of these species of Moorella display a similar substrate range. Both species also require the vitamin nicotinic acid for growth (Lundie and Drake 1984; Savage and Drake 1986). The substrate range of M. thermoautotrophica includes H2-CO2, CO, formate, methanol, glucose, fructose, glycerate, glycolate, and methoxylated aromatic compounds (Wiegel et al. 1981; Fro¨stl et al. 1996; Seifritz et al. 1999). Nitrate is utilized as an alternative electron acceptor and is dissimilated to nitrite and ammonium;

Acetogenic Prokaryotes

ethanol and n-propanol are growth-supportive substrates only when nitrate is available for dissimilation (Fro¨stl et al. 1996). Natroniella acetigena. This organism is a haloalkaliphilic acetogen and was isolated from the soda deposits at Lake Magadi, Kenya (Zhilina et al. 1996). Cells are Gram-negative, sporeforming, motile, large rods. The substrate range is limited and includes lactate, pyruvate, ethanol, glutamate, and propanol. Growth does not occur on H2-CO2 or CO-CO2. Acetate is the sole reduced end product. Propionate is formed during growth on propanol. The optimal pH is 10, and the optimal salinity for growth is 12 % NaCl (w/v). Natronincola histidinovorans. This species is a moderately haloalkaliphilic acetogen and was isolated from soda deposits at Lake Magadi, Kenya (Zhilina et al. 1998). Cells are Grampositive, motile rods; sporeforming and nonsporeforming strains have been isolated. Natronincola histidinovorans is specialized in using amino acids (histidine, glutamate, and casaminoacids) as sources of energy. Neither H2-CO2 nor CO-CO2 support growth. Optimal growth occurs at pH 9 and a salinity of 9 % NaCl. Acetate and ammonium are the main end products. Oxobacter pfennigii. This organism was isolated from the rumen fluid of a steer and was first described as Clostridium pfennigii (Krumholz and Bryant 1985). On the basis of phylogenetic analysis of the 16S rRNA gene sequence, C. pfennigii was reclassified as O. pfennigii (Collins et al. 1994). Cells are Grampositive, motile, sporeforming, slightly curved rods. Substrates include CO, pyruvate, vanillate, vanillin, ferulate, syringate, and trimethoxybenzoate. In contrast to most other acetogens, acetate is not produced from methoxybenzenoids (O-methyl groups are utilized, and butyrate and the respective hydroxybenzenoids are formed; Krumholz and Bryant 1985). During growth on CO or pyruvate, acetate is formed in addition to butyrate or is the sole product, respectively. Ruminococcus hydrogenotrophicus. This species is a nonsporeforming coccobacillus that was isolated from human feces (Bernalier et al. 1996c). Ruminococcus hydrogenotrophicus grows on H2-CO2, formate, pyruvate, and several sugars. Acetate is the sole product from H2-CO2-dependent growth; however, glucose and fructose are metabolized to acetate, lactate, ethanol, and small amounts of isobutyrate and isovalerate (Bernalier et al. 1996c). Thus, the metabolism of sugars involves several fermentative processes. Ruminococcus productus. This organism was originally isolated from various mammalian gastrointestinal tracts and was described as Peptostreptococcus productus; the original isolates were not described as acetogens (Moore and Holdeman 1974; Varel et al. 1974; Holdeman-Moore et al. 1986). On the basis of phylogenetic analysis of the 16S rRNA gene sequence, P. productus was reclassified as R. productus (Ezaki et al. 1994). Two acetogenic strains (strain U-1 [ATCC 35244] and strain Marburg [ATCC 43917]) of R. productus have been isolated from sewage sludge (Lorowitz and Bryant 1984; Geerligs et al. 1987). Cells are Gram-positive, nonsporeforming, nonmotile elongated cocci occuring often in pairs or chains (Lorowitz and Bryant 1984; Holdeman-Moore et al. 1986; Geerligs et al. 1987). Growth-supportive substrates of the acetogenic strains

1

include CO, H2-CO2, monomeric and dimeric sugars, and methoxylated aromatic compounds; growth is particularly good on CO (Lorowitz and Bryant 1984; Geerligs et al. 1987; Parekh et al. 1992). The acrylate side chain of methoxylated and nonmethoxylated phenylacrylates can be used as alternative electron acceptor (Parekh et al. 1992; Misoph et al. 1996b). The major reduced end product is acetate; however, under CO2-limited conditions or when substrate concentrations are high (e.g., 10 mM fructose), lactate, succinate, and formate are also formed (Misoph and Drake 1996a). Ruminococcus schinkii. This organism was isolated from rumen content of 1–3-day-old lambs (Rieu-Lesme et al. 1996b). Cells are Gram-positive, nonsporeforming, nonmotile cocci. Substrates include H2-CO2, various sugars, glycerol, syringate, and ferulate. Acetate is the sole reduced end product. Sporomusa acidovorans. This species was isolated from a distillation wastewater fermentor (Ollivier et al. 1985a). Cells are Gram-negative, sporeforming, motile, curved rods. Growthsupportive substrates mainly include organic acids, H2-CO2, methanol, glycerol, and a few sugars; acetate is the sole reduced end product with all substrates. Sporomusa aerivorans. This organism was isolated from a soil-feeding termite (Boga et al. 2003). Cells are Gram-negative, sporeforming, motile, curved rods. Growth-supportive substrates include H2-CO2, formate, methanol, ethanol, lactate, pyruvate, mannitol, citrate, and various methoxylated aromatic compounds; hexoses are not utilized. Cells tolerate and consume small amounts of oxygen and are catalase positive (Boga and Brune 2003). Sporomusa malonica. This species was isolated from freshwater sediment (Dehning et al. 1989). Cells are Gram-negative, sporeforming, motile, curved rods. The organism exhibits a very versatile metabolism and utilizes H2-CO2 and numerous organic compounds, including formate, pyruvate, alcohols, dicarboxylic acids, fructose, and trimethoxycinnamate. Acetate is the reduced end product when typical acetogenic substrates such as H2-CO2, formate, methanol, fructose, pyruvate, or the O-methyl groups of trimethoxycinnamate are metabolized (Dehning et al. 1989). Alcohols yield acetate and the respective fatty acids, and crotonate and 3-hydroxybutyrate yield acetate and butyrate. As with relatively few anaerobes, S. malonica metabolizes simple dicarboxylic acids (e.g., malonate and succinate) by decarboxylation to the respective fatty acids. Sporomusa ovata. This organism was isolated from sugar beet leaf silage (Mo¨ller et al. 1984). Cells are Gram-negative, sporeforming, motile, curved rods. Growth is supported by a variety of substrates including H2-CO2, pyruvate, lactate, alcohols, fructose, betaine, dimethylglycine, and sarcosine. Acetate is the sole reduced end product; methylamines are formed from N-methyl compounds. Reductively dechlorinates tetrachloroethylene to trichloroethylene (Terzenbach and Blaut 1994). Cultures demethylate the osmolytes dimethylsulfoniopropionate and glycine-betaine to methylthiopropionate and dimethylglycine, respectively; however, only the demethylation of glycine-betaine supports growth of the organism (Jansen and Hansen 2001).

19

20

1

Acetogenic Prokaryotes

. Fig. 1.7 Electron micrograph of a vegetative cell of Sporomusa silvacetica (DSM 10669T) showing flagella inserting at the concave side of the cell [From Kuhner et al. (1997), used with permission from International Union of Microbiological Societies]

Sporomusa paucivorans. This species was isolated from lake sediment (Hermann et al. 1987). Cells are Gram-negative, nonsporeforming, motile, slightly curved rods. H2-CO2, formate, methanol, pyruvate, serine, betaine, alcohols, and ethylene glycol support growth. Acetate is the sole reduced end product. Oxidation of alcohols yields the corresponding fatty acids. Sugars are not utilized. Sporomusa silvacetica. This organism was isolated from forest soil (Kuhner et al. 1997). Cells are Gram-negative, sporeforming, motile, slightly curved rods (> Fig. 1.7). Growth occurs on H2-CO2, formate, methanol, pyruvate, vanillate, ferulate, fructose, betaine, fumarate, 2,3-butanediol, ethanol, lactate, and glycerol. With most substrates, acetate is the main reduced end product. Fumarate is dismutated to acetate and succinate. Vanillate and ferulate are O-demethylated and reduced, respectively. Cells tolerate and consume small amounts of oxygen (Karnholz et al. 2002). Sporomusa sphaeroides. This species was isolated from river mud (Mo¨ller et al. 1984). Cells are Gram-negative, sporeforming, motile, curved rods. Growth occurs on H2-CO2, pyruvate, lactate, alcohols, glycerol, serine, ethyleneglycol, betaine, and other N-methyl compounds. Acetate is the sole reduced end product; methylamines are formed from N-methyl compounds. Cultures demethylate the osmolytes dimethylsulfoniopropionate and glycine-betaine to methylthiopropionate and

dimethylglycine, respectively; however, only the demethylation of glycine-betaine supports growth of the organism (Jansen and Hansen 2001). Sporomusa termitida. This organism was isolated from the gut of a wood-feeding termite (Breznak et al. 1988). Cells of S. termitida are Gram-negative, sporeforming, motile, straight to slightly curved rods. Substrates include H2-CO2, CO, formate, methanol, ethanol, betaine, sarcosine, lactate, pyruvate, oxaloacetate, citrate, malonate, succinate, mannitol, and trimethoxybenzoate. Acetate is the main reduced end product. As with S. malonica, S. termitida decarboxylates succinate to propionate (Breznak et al. 1988; Dehning et al. 1989). Sporomusa termitida grows mixotrophically, e.g., by utilizing H2 and methanol or lactate at the same time (Breznak and Switzer Blum 1991). Syntrophococcus sucromutans. This organism is a Gramnegative, nonsporeforming, nonmotile, coccoid bacterium that was isolated as a dominant methoxybenzenoids utilizer from the rumen contents of a steer (Krumholz and Bryant 1986). Syntrophococcus sucromutans has a unique metabolism: growth with carbohydrates or pyruvate is only possible in the presence of electron acceptors such as formate, O-methyl groups, or a hydrogenotrophic methanogen (Krumholz and Bryant 1986). Formate and O-methyl groups are metabolized via an acetyl-CoA pathway that lacks formate dehydrogenase and is therefore incomplete (Dore´ and Bryant 1990). Thermoacetogenium phaeum. This species is a thermophilic acetogen that was isolated from an anoxic pulp wastewater reactor (Hattori et al. 2000). Cells are Gram-positive, sporeforming, nonmotile, straight or slightly curved rods. Substrates include H2-CO2, formate, methanol, n-propanol, methoxylated benzoic acids, glycine, and cysteine. Acetate is the sole reduced end product. Acetate is oxidized in the presence of hydrogenotrophic methanogens or an alternative electron acceptor (e.g., sulfate or thiosulfate); concomitantly, methane is produced by the syntrophic methanogen or the alternative electron acceptor is reduced. Its ability to oxidize acetate in syntrophic association with hydrogenotrophic methanogens is similar to that of two other anaerobic acetate oxidizers, strain AOR and Clostridium ultunense (Zinder and Koch 1984; Lee and Zinder 1988; Schnu¨rer et al. 1996). Thermoanaerobacter kivui. This species is a thermophilic acetogen that was isolated from lake sediments of Lake Kivu, Africa, and was first described as Acetogenium kivui (Leigh et al. 1981). On the basis of phylogenetic analysis of the 16S rRNA gene sequence, A. kivui was reclassified as T. kivui (Rainey et al. 1993; Collins et al. 1994). Cells are nonmotile, nonsporeforming rods often occurring in pairs or chains (Leigh et al. 1981; > Fig. 1.8). The cell wall is covered by a hexagonally structured S-layer consisting of an 80-kDa protein (Rasch et al. 1984; Lupas et al. 1994). The temperature optimum is 66  C. Autotrophic growth occurs on H2-CO2, and heterotrophic growth occurs on glucose, mannose, fructose, pyruvate, and formate; acetate is the main reduced end product (Leigh et al. 1981). Growth does not occur on CO-CO2 (Daniel et al. 1990). Thermoanaerobacter kivui grows robustly on H2-CO2, a substrate that yields very

Acetogenic Prokaryotes

. Fig. 1.8 Phase-contrast photomicrograph of cells of Thermoanaerobacter kivui. Bar equals 5 mm [From Leigh et al. (1981), used with permission from Springer. The micrograph was kindly provided by R.S. Wolfe]

slow, poor growth with most acetogens, and displays exceptionally high specific activities of hydrogenase and CO dehydrogenase (i.e., acetyl-CoA synthase) when cultivated chemolithoautotrophically on H2-CO2 (Daniel et al. 1990). Cells tolerate and consume small amounts of oxygen (Karnholz et al. 2002). Treponema primitia. This acetogenic spirochete was isolated from the hindgut of termites (Graber et al. 2004b; Graber and Breznak 2004a). Growth occurs on H2-CO2, certain mono- and disaccharides, and methoxybenzoids. It can use H2-CO2 and organic compounds simultaneously, requires folate for growth, and can tolerate low amounts of O2; cells have NADH peroxidase and NADH oxidase activities.

Cultivation Methods Even though many acetogens likely have the ability to tolerate small amounts of O2 (Ku¨sel et al. 2001; Karnholz et al. 2002; Boga and Brune 2003), acetogens should be considered obligate anaerobes, and care should be taken in the laboratory to protect them from oxic conditions. The Hungate technique (Hungate 1969) or modifications thereof are recommended for cultivation purposes. Growth media should be anoxic; sodium sulfide, cysteine, dithionite, or dithiothreitol are often used in cultivation media. Titanium (III) reducer has also been used; it may be

1

less toxic than sulfide-based reducers (Zehnder and Wuhrmann 1976; Moench and Zeikus 1983). Sulfide-based reducers can decrease the number of acetogens obtained from aerated soils (Ku¨sel et al. 1999c). Cadmium chloride (CdCl2) has also been used as reducing agent for acetogen media (Breznak and Switzer 1986; Breznak et al. 1988). > Table 1.3 outlines the contents of a medium that can be used for the cultivation of most known acetogens. In general, acetogens prefer near neutral to slightly alkaline pH. However, many decades ago, Wieringa (1941) reported that enrichment of acetogens under alkaline conditions (pH 8–9) might favor their isolation. It is thus noteworthy that the first haloalkaliphilic acetogens, N. histidinovorans and N. acetigena, were recently isolated from soda-depositing lakes; these acetogens only grow between pH 8 and 11 (Zhilina et al. 1996, 1998). Obviously, designing media for the cultivation of acetogens must take into consideration the in situ conditions of the habit under investigation. Since CO2 is used as a terminal electron acceptor by acetogens, many acetogens cannot grow under certain conditions unless CO2 is readily available (see the section on > ‘‘CO2 as Terminal Electron Acceptor and the Concept of Fermentation’’ in this chapter). Thus, acetogenesis is optimized in acetogen media containing a source of CO2. Although some chemolithoautotrophic acetogens grow without any trace organic nutrients (e.g., T. kivui; Leigh et al. 1981), many acetogens require supplemental vitamins. The inclusion of trace metals in acetogen media is important because acetogens are rich in metalloenzymes (Ljungdahl 1986). Many acetogens have unknown nutritional factors. For example, the protocol used to elucidate the nutritional requirements of M. thermoacetica (Lundie and Drake 1984) and M. thermoautotrophica (Savage and Drake 1986) was not successfully applied to R. productus (supplemental vitamins, amino acids, fatty acids, and other nutrients did not substitute for unknown growth factors in yeast extract; thus, the nutritional requirements for R. productus remain unresolved; H. L. Drake and coworkers, unpublished data). The substrate used to enrich and isolate an acetogen can be selective for a certain catabolic type. H2-CO2 is quite often selective for acetogens and has been used for the isolation of numerous acetogens [e.g., T. kivui (Leigh et al. 1981), A. woodii (Balch et al. 1977), and S. termitida (Breznak et al. 1988)]. CO-CO2 is also selective (e.g., R. productus; Lorowitz and Bryant 1984; Geerligs et al. 1987). The capacity to utilize methoxylated aromatic compounds is a widespread catabolic potential of acetogens; thus, methoxylated aromatic compounds can also be used to selectively enrich and isolate acetogens (e.g., A. woodii; Bache and Pfennig 1981). Many isolates have been enriched and isolated with compounds not typically utilized by anaerobes. Examples of such substrates include mandelate (Acetobacterium strain AmMan1; Do¨rner and Schink 1991), trimethylamine (A. arabaticum; Zhilina and Zavarzin 1990), methoxyacetate (strain RMMac1; Schuppert and Schink 1990), and methyl chloride (‘‘Acetobacterium dehalogenans’’ [formerly named ‘‘strain MC’’; Traunecker et al. 1991]). Isolation can be accomplished by various methods, such as the roll-tube method of

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22

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Acetogenic Prokaryotes

. Table 1.3 Contents of a typical acetogen mediuma Salts

mg/l

NaCl

400

NH4Cl

400

MgCl2·6H2O

330

CaCl2·2 H2O

150

Trace elements MnSO4·H2O

mg/l 2.5

FeSO4·7H2O

0.5

Co(NO3)2·6H2O

0.5

ZnCl2

0.5

NiCl2·6H2O

0.25

H2SeO4

0.25

CuSO4·5H2O

0.05

AlK(SO4)2·12H2O

0.05

H3BO3

0.05

Na2MoO4·2H2O

0.05

Na2WO4·2H2O

0.05

Reducers

mg/l

Cysteine·HCl·H2O

250

Sodium sulfide

250

Vitamins

mg/l

Nicotinic acid

0.25

Cyanocobalamin

0.25

p-Aminobenzoic acid

0.25

Calcium D-pantothenate

0.25

Thiamine·HCl

0.25

Riboflavin

0.25

Lipoic acid

0.30

Folic acid

0.1

Biotin

0.1

Pyridoxal HCl

0.05

Buffer for pH 6.7 media

mg/l

NaHCO3

7,500

KH2PO4

500

Gas phase

100 % CO2

Buffer for pH 7.8 media

mg/l

NaCO3

16,000

Na2CO3·10H2O

16,500

KH2PO4

1500

Gas phase

100 % N2

a Media should be prepared using anoxic techniques, and resazurin (1 mg/l) can be used as a redox indicator. Media can be enriched with yeast extract (0.5–2 g/l), tryptone (0.5–2 g/l), clarified rumen fluid (50 ml/l), or casamino acids (0.5–2 g/l). Crimp-sealed tubes or bottles are recommended for cultivation. Compiled from Daniel et al. (1990) and Matthies et al. (1993)

Hungate (1969), agar shake dilution series (Pfennig 1978), or streaking/plating on agar or Gelrite® (preferred for thermophiles because of its increased capacity to remain solid at higher temperatures). Methanogens can usually be excluded during enrichment and isolation with 10–50 mM bromoethanesulfonate (an analog of coenzyme M; Gunsalus et al. 1978; Zehnder et al. 1980; Smith and Mah 1981; Greening and Leedle 1989; Kane and Breznak 1991a). Lumazine (2,4-[1H,3H]-pteridinedione) is also an inhibitor of methanogens (Methanobacterium thermoautotrophicum does not grow in media containing 0.1 mM lumazine; Nagaranthal and Nagle 1992) and might also be useful in such enrichments. Lumazine appears to inhibit methyl-S-CoM reductase (Nagaranthal and Nagle 1992). Likewise, N-substituted derivatives of p-aminobenzoic acid inhibit the enzyme responsible for the synthesis of methanopterin (an intermediate in methanogenesis) and can block the growth of certain methanogens but do not interfere with the growth of acetogens (Dumitru et al. 2003).

Taxonomy and Phylogeny Acetogens do not form a taxonomic group of closely related organisms. Although utilization of the acetyl-CoA pathway unifies them, they are extremely diverse genetically; the G + C content of their genomes varies from 22 mol% for C. ljungdahlii to 62 mol% for H. foetida (> Table 1.2). However, all acetogenic isolates to date are members of the domain bacteria, including the extreme halophilic and haloalkaliphilic isolates A. arabaticum and N. acetigena, respectively (Zhilina and Zavarzin 1990; Zhilina et al. 1996). The phylogeny of most acetogens with validated genus and species names and a couple of other acetogens not yet included in the ‘‘List of Bacterial Names with Standing in Nomenclature’’ ([{http://www.bacterio.cict.fr}]) has been determined in either comprehensive analyses of bacteria (e.g., within a certain bacterial group, such as the clostridia) or in the first description of a respective species. Analyses of phylogenetic relationships resulted in the reclassification of several acetogenic species (> Table 1.4). Within the phylogenetic tree of the bacteria, acetogens are found in more than one phylum and do not form tight clusters like methanogens or sulfate-reducing bacteria. However, most acetogens are affiliated with the phylum of the Gram-positive bacteria with low DNA G + C content (> Fig. 1.9). One genus that harbors acetogenic species, Treponema (> Fig. 1.5), is affiliated with the phylum Spirochetes. Holophaga, a genus with only one species isolated to date, H. foetida, represents a new line of descent; together with the nonacetogen Geothrix fermentans, H. foetida was one of the two founding species of the phylum Holophaga/Acidobacterium (Liesack et al. 1994; Ludwig et al. 1997; Leadbetter et al. 1999; Lilburn et al. 1999). The 16S rRNA genes of the species belonging to the genus Acetoanaerobium have not been sequenced; thus, the phylogenetic positions of the acetogens A. notorae and ‘‘A. romashkovii’’

Acetogenic Prokaryotes

. Table 1.4 Acetogens that have been reclassified Described as

Reclassified as

Reference

Acetogenium kivui

Thermoanaerobacter Collins et al. (1994) kivui

Clostridium fervidusa

Caloramator fervidus Collins et al. (1994)

Clostridium thermoaceticum

Moorella thermoacetica

Collins et al. (1994)

Clostridium thermoautotrophicum

Moorella thermoautotrophica

Collins et al. (1994)

Clostridium pfennigii

Oxobacter pfennigii

Collins et al. (1994)

Peptostreptococcus productus

Ruminococcus productus

Ezaki et al. (1994)

a

It is not certain that this organism is an acetogen

are unclear. On the basis of morphological and physiological properties, A. notorae has been tentatively assigned to the Bacteroidaceae (Sleat et al. 1985). Acetogens affiliate with 8 (I, V, VI, IX, XI, XII, XIVa, XV) out of the 19 Clostridium clusters within the phylum of the Grampositive bacteria with low DNA G + C content (Collins et al. 1994; Zhilina et al. 1998; Stackebrandt et al. 1999; > Fig. 1.9). Although some genera (e.g., Acetobacterium, Sporomusa, and Moorella) are composed exclusively of acetogenic species, many acetogens in these phylogenetic clusters are very closely related to nonacetogenic bacteria (> Fig. 1.10). Indeed, genera often include acetogenic and nonacetogenic species (e.g., Clostridium, Ruminococcus, Eubacterium, and Thermoanaerobacter). For example, the closest relative of C. formicoaceticum is the nonacetogen Clostridium felsineum (99.3 % sequence similarity). In some cases, clostridial species were not originally described as acetogens but were later discovered to possess acetogenic capabilities. For example, C. coccoides strain 1410 and C. scatologenes strain SL1 were isolated as acetogens, and subsequent physiological studies revealed that the type strains of these two species also have acetogenic capabilities (Kamlage et al. 1997; Ku¨sel et al. 2000). The opposite is also true, as seen in the case of C. glycolicum. Two strains of C. glycolicum, strain RD-1 and strain 22, are acetogens, yet the type strain of C. glycolicum does not grow acetogenically (Ohwaki and Hungate 1977; Ku¨sel et al. 2001). Thus, certain acetogens might lose their acetogenic capabilities after prolonged laboratory cultivation. An acetogenic strain of C. difficile (AA1) has been isolated from the rumen contents of a newborn lamb; whether the pathogenic type strain of C. difficile has acetogenic capabilities is unknown (Hall and O’Toole 1935; Cato et al. 1986; Rieu-Lesme et al. 1998).

Detection of Acetogens On agar or Gelrite®, acetogenic colonies can be differentiated from nonacetogenic colonies by their ability to form (1) clear

1

zones on media containing calcium carbonate (Balch et al. 1977) or (2) colored zones on media supplemented with a pH indicator (e.g., bromocresol green; Braun et al. 1979; Leedle and Greening 1988). A colorimetric most-probable-number assay can be utilized to enumerate acetogens that are capable of O-demethylating methoxylated lignin derivatives (e.g., vanillate; Harriott and Frazer 1997). This method based on a yellowing colorimetric reaction between Ti(III) and the vicinal hydroxyl groups of the O-demethylated aromatic product (Moench and Zeikus 1983; Kreft and Schink 1993; Harriott and Frazer 1997) has been used to detect acetogens on the roots of the sea grass Halodule wrightii (Ku¨sel et al. 1999b). The occurrence of acetyl-CoA synthase in an isolate can be considered good evidence that the organism is an acetogen. As outlined in the section on > ‘‘Enzymology of the Acetyl-CoA Pathway,’’ this enzyme catalyzes two reactions that can be assayed. One of these reactions, i.e., that of CO dehydrogenase, is inexpensive and easily measured by monitoring the reduction of an artificial electron carrier (e.g., methyl viologen) with a spectrophotometer. Thus, CO dehydrogenase activity is often used to assess acetyl-CoA synthase. However, the occurrence of CO dehydrogenase activity can be misleading and cannot be taken as definitive evidence that an organism utilizes the acetyl-CoA pathway. For example, the CO dehydrogenase activity of Clostridium pasteurianum (Fuchs et al. 1974; Thauer et al. 1974; Diekert and Thauer 1978) implies that this bacterium is an acetogen, but it is not. It is therefore essential that additional evidence (e.g., precise substrate/product stoichiometries or additional enzymological information) be obtained before concluding that an organism is an acetogen (i.e., utilizes the acetylCoA pathway). Although the acetyl-CoA synthase assay is more difficult than that for CO dehydrogenase, acetyl-CoA synthase activity can be assayed with the pyruvate/CO-coupled methyltetrahydrofolate assay (Schulman et al. 1973; Drake et al. 1981a; Hu et al. 1982). This activity is much more conclusive relative to the acetogenic nature of an organism. The acetyl-CoA synthase assay is classically based on the use of [14CH3]methyltetrahydrofolate and the measurement of 14C-labeled acetate (which is derived from the 14C-labeled acetyl-CoA that is formed by acetyl-CoA synthase). However, a method that does not involve the use of 14C can also be used; in this assay, the acetyl-CoA formed by acetyl-CoA synthase is converted to acetate, which is then measured by high-performance liquid chromatography (Fro¨stl et al. 1996). The activities obtained with the CO dehydrogenase and acetyl-CoA synthase activity assays do not always yield parallel results, and the two respective activities can vary differentially (Kellum and Drake 1986). In general, caution is needed when drawing conclusions based on results obtained with these assays. The apparent importance of acetogens to the overall flow of carbon, reductant, and energy in many habitats has generated an interest in assessing acetogenic populations in environmental samples with molecular methods. Unfortunately, based on their 16S rRNA gene sequences, acetogens are not a monophyletic

23

Low G+C Gram-positive Bacteria

. Fig. 1.9 Parsimony tree containing 20 genera of acetogenic bacteria; tree is based on full length 16S rRNA sequences (ARB-release June 2002). The genus Acetoanaerobium (> Table 1.2) is not represented in the tree because a sequence for a species of this genus is not available. Comparison of the tree topology with maximum-likelihood and neighbor-joining-based trees suggests that the phylogeny of acetogens remains to be resolved. Numbers are sequence accession numbers. Bar corresponds to 10 nucleotide substitutions per 100 sequence positions. Though unavailable when the tree was constructed, please note that the two Treponema strains have been recently named ‘‘T. primitia’’ (Graber et al. 2004b)

Acetitomaculum ruminis, M59083 Syntrophococcus sucromutans, Y18191 “Bryantella formatexigens”, AJ318527 Clostridiummethoxybenzovorans, AF067965 Ruminococcus schinkii, X94965 Ruminococcus hydrogenotrophicus, X95624 Ruminococcus productus, X94966 Clostridium coccoides, M59090 Clostridium coccoides 1410, Y10584 Clostridium difficile AA1, AJ310756 Clostridium glycolicum, AJ291746 Clostridium mayombei, M62421 Clostridium formicoaceticum, X77836 Clostridium aceticum, Y18183 Natronincola histidinovorans, Y16716 Clostridium ultunense, Z69293 Acetobacterium tundrae, AJ297449 Acetobacterium paludosum, X96958 Acetobacterium bakii, X96960 Acetobacterium fimetarium, X96959 Acetobacterium carbinolicum, X96956 “Acetobacterium psammolithicum”, AF132739 Acetobacterium malicum, X96957 Acetobacterium woodii, X96954 Acetobacterium wieringae, X96955 “Butyribacterium methyiotrophicum”, AF064241 Eubacterium limosum, M59120 Eubacterium aggregans, AF073898 “Clostridium autoethanogenum”, Y18178 Clostridium ljungdahlii, M59097 Clostridium scatologenes SL1, Y18813 Clostridium scatologenes, M59104 Clostridium magnum, X77835 Oxobacter pfennigii, X77838 Caloramator fervidus, L09187 Thermoanaerobacter kivui, L09160 Thermoacetogenium phaeum, AB020336 Moorella thermoautotrophica, L09168 Moorella thermoacetica, M59121 0.10 “Moorella mulderl”, AF487538 Moorella qlycerini, U82327

Holophaga/Acidobacterium

Spirochetes

1

Sporomusa silvacetica, Y09976 Sporomusa ovata, AJ279800 Sporomusa acidovorans, AJ279798 Sporomusa malonica, AJ279799 Sporomusa paucivorans, M59117 Sporomusa sphaeroides, AJ279801 Sporomusa termitida, M61920 Sporomusa aerivorans, AJ506191 Acetonema longum, AJ010964 Natroniella acetigena, X95817 Acetohalobium arabaticum, L37422

Holophaga foetida, X77215

Treponema ZAS-1, AF093251

Treponema ZAS-2, AF093252

24 Acetogenic Prokaryotes

Acetogenic Prokaryotes

1

Hologpaga foetida “Geothrix fermentans” Treponema sp. ZAS-1 Thermophilic spirochete Orenia marismortui Natroniella acetigena Acetohalobium arabaticum Sporohalobacter lortetii Syntrophococcus sucromutans Eubacterium cellulosolvens “Clostridium methoxybenzovorans” Clostridium indolis Clostridium coccoides Clostridium coccoides 1410 Ruminococcus obeum Ruminococcus schinkii Ruminococcus hansenii Ruminococcus hydrogenotrophicus Eubacterium contortum Clostridium polysaccharolyticum Acetitomacutum ruminis Clostridium glycolicum Clostridium glycolicum RD-1 Clostridium felsineum Clostridium formicoaceticum Clostridium aceticum Clostridium hastiforme Clostridium ultunense Acetobacterium woodii Eubacterium callanderi Eubacterium limosum Eubacterium barkeri Eubacterium aggregans Clostridium ljungdahlii Clostridium tyrobutyricum Clostridium scatologenes SL1 Clostridium scatologenes Clostridium tetanomorphum Caloramator indicus Caloramator fervidus Sporomusa sphaeroides Acetonema longum Dendrosporobacter quercicolus Thermoanaerobacter kivui Thermoanaerobacter brockii Thermoanaerobacter ethanolicus Moorella thermoacetica Moorella glycerini Thermoterrabacterium ferrireducens Thermoacetogenium phaeum 0.10

. Fig. 1.10 Parsimony tree of selected acetogenic bacteria (blue, bold font) and their closest nonacetogenic relatives (red, nonbold font) based on full length 16S rRNA sequences (ARB-release December 1998). Bar corresponds to 10 nucleotide substitutions per 100 sequence positions

group, and many acetogens are very closely related to nonacetogenic species (see the section on > ‘‘Taxonomy and Phylogeny’’ in this chapter). Thus, the development of 16S rRNA oligonucleotide probes and primers that exclusively target all known acetogens is likely an impossible task. Nonetheless, a 16S rRNA oligonucleotide probe (probe AW) has been developed that targets the acetogenic species that form a phylogenetically tight cluster in the genus Acetobacterium; this probe is also specific for the acetogen E. limosum (Ku¨sel

et al. 1999b). Another 16S rRNA oligonucleotide probe (probe Clost I) is specific for a few acetogens and many nonacetogens in clostridial clusters I and II (Ku¨sel et al. 1999b). Oligonucleotide probes or primers that target functional genes can also be used to assess microorganisms in environmental samples, and this approach has been used to evaluate several microbial groups, including ammonia-oxidizing bacteria, denitrifying bacteria, and nitrogen-fixing bacteria (Rotthauwe et al. 1997; Braker et al. 2000; Lovell et al. 2000).

25

26

1

Acetogenic Prokaryotes

Formyltetrahydrofolate synthetase (FTHFS) is one of the key enzymes in the acetyl-CoA pathway and catalyzes the ATPdependent synthesis of formyltetrahydrofolate from formate and tetrahydrofolate. FTHFS is highly conserved in acetogens. A FTHFS-based functional group-specific DNA probe and FTHFS-based primers for polymerase chain reaction (PCR) amplification of partial FTHFS gene sequences have been utilized for evaluating the occurrence of acetogens in environmental samples and complex natural populations (Lovell and Hui 1991; Lovell 1994; Leaphart and Lovell 2001; Leaphart et al. 2003; Salmassi and Leadbetter 2003).

The Acetyl-CoA Pathway and Bioenergetics The acetyl-CoA pathway is composed of two reductive branches (i.e., the methyl branch and the carbonyl branch), both of which reduce CO2 and fix CO2-derived carbon into covalent bond forms (> Fig. 1.11). The acetyl-CoA pathway can be presented in a cyclic form (e.g., Wood and Ljungdahl 1991; Ragsdale 1997); however, it is a linear process that does not depend on multicarbon intermediates to which CO2 is fixed in a cyclic fashion (e.g., the Calvin cycle and the reductive tricarboxylic acid cycle are cyclic, CO2-fixing processes that are dependent upon ribulose biphosphate and oxalacetate, respectively, for the initial fixation of CO2). Although the cofactors and electron carriers of the pathway cycle between different states, the pathway itself is linear relative to carbon flow (> Fig. 1.11). The acetyl-CoA pathway is a terminal-electron-accepting process that also provides a mechanism for the assimilation of CO2 and other sources of carbon into biomass (Ljungdahl and Wood 1965; Eden and Fuchs 1982, 1983; Ljungdahl 1986). The main function of the pathway may vary with the growth conditions of the cell. The biochemistry of the acetyl-CoA pathway has been described in numerous reviews (Ljungdahl 1986; Ragsdale 1991, 1994, 1997, 2004; 1991a; Wood and Ljungdahl 1991; Diekert and Wohlfarth 1994a, 1994b; Drake 1994; Ragsdale and Kumar 1996; Drake et al. 1997; Das and Ljungdahl 2000; Drake and Ku¨sel 2003); many of these references also provide historical perspectives on the studies that resolved the enzymological features of the pathway. A gateway to some of the more recent studies on the enzymology of the acetyl-CoA pathway can be found in the following references: Maynard and Lindahl (1999, 2001), Furdui and Ragsdale (2000), Radfar et al. (2000), Ragsdale (2000, 2003a, b, 2004), Mu¨ller et al. (2001, 2004), Leaphart et al. (2002), Lindahl (2002), Banerjee and Ragsdale (2003), Das and Ljungdahl (2003), Bramlett et al. (2003), Darnault et al. (2003), Grahame (2003), and Loke and Lindahl (2003). Acetyl-CoA synthase is a centrally important enzyme of the acetyl-CoA pathway, and many prokaryotes make use of enzymes that are closely related to the acetyl-CoA synthase of acetogens (see the sections on > ‘‘Enzymology of the AcetylCoA Pathway’’ and > ‘‘Occurrence of the Acetyl-CoA Pathway in Nonacetogenic Microorganisms’’ in this chapter). Numerous

theortetical considerations suggest that an acetyl-CoA-synthasedependent pathway may have constituted the first autotrophic process on earth (Fuchs 1986; Wood and Ljungdahl 1991; Lindahl and Chang 2001). It is uncertain whether the pathway first evolved for the purpose of carbon assimilation (i.e., the reduction and fixation of CO2) or the oxidation of acetate. Phylogenetic evaluations of acetyl-CoA synthases indicate that microorganisms (e.g., acetogens and methanogens) that have this enzyme, or enzymes that are closely related to it, had a common ancestor (Lindahl and Chang 2001). This section will focus on some of the biochemical and enzymological features of the acetyl-CoA pathway of acetogens.

CO2 as Terminal Electron Acceptor and the Concept of Fermentation The main function of the acetyl-CoA pathway during growth on sugars is the recycling of reduced electron carriers (NAD, ferredoxin, etc.; > Fig. 1.12). The eight reducing equivalents that are collectively generated during glycolysis and the oxidation of pyruvate are used to reduce CO2 to acetate via the acetyl-CoA pathway. During growth on glucose, the cell has ready access to ATP formed via substrate-level phosphorylation and to biosynthetic precursors (via the breakdown of glucose). Thus, the lithoautotrophic functions of the pathway (i.e., the conservation of energy and the production of acetyl-CoA for anabolism and the assimilation of carbon) are probably of minor importance under such conditions. CO2 is the terminal electron acceptor of acetogens when they are grown under acetogenic conditions (see the section on > ‘‘Use of Diverse Terminal Electron Acceptors’’ in this chapter); it is therefore important to include an adequate supply of CO2 (or carbonates) in the growth medium when acetogens are cultivated in the laboratory (see the section on > ‘‘Cultivation Methods’’ in this chapter). This point may seem obvious, but it is not generally appreciated why supplemental CO2 is essential to the growth of acetogens, since the stoichiometries for the synthesis of acetate from numerous substrates do not indicate that supplemental CO2 is required. For example, even though the stoichiometric conversion of sugars (e.g., glucose or fructose) to acetate (reaction > 1.2 above) indicates that supplemental CO2 is not required for acetogenesis, growth on sugars, as well as the metabolism of sugars, may be significantly impaired in the absence of supplemental CO2 (Andreesen et al. 1970; O’Brien and Ljungdahl 1972; Braun and Gottschalk 1981). The reason why exogenous CO2 is required for optimal growth on glucose is illustrated in > Fig. 1.13. Supplemental CO2 is required for the recycling of reduced electron carriers, and this intracellular management of reductant must occur before glucose-derived CO2 becomes available via the decarboxylation of pyruvate. In support of the scheme illustrated in 14 > Fig. 1.12, early C studies demonstrated that carbons 3 and 4 of glucose mostly enter the pool of CO2 rather than acetate (Wood 1952b; O’Brien and Ljungdahl 1972). Thus,

Acetogenic Prokaryotes

Methyl Branch

Carbonyl Branch

CO2 +3 [+22]

1

CO2

2 e-

(4) HCOOH

−8

ATP (5) (11)

ADP + Pi +

[CHO]-THF

H+ −4 (6, 7)

H2O

2 e-

[CH]-THF 2 e-

−23 [−5] (6, 7)

+20 [+38]

[CH2]-THF 2 e-

−40 [−22] (8)

Energy Conserved

[CH3]-THF

−38 (10−12)

(9) [CH3]-[Co-Enzyme]

[CO]

HSCoA O C a t a b o l i s m

CH3C-SCoA +9 (2)

Pi

CH3COO-PO32− ADP

−13 (3)

ATP CH3COOH −1

(Overall: −95kJ mol )

Cell Carbon

A n a b o l i s m

. Fig. 1.11 The acetyl-CoA ‘‘Wood/Ljungdahl’’ pathway. The numbers (in black) adjacent to the reactions are standard Gibbs free energies in kJ mol 1 (values have been rounded off). For the four reactions in which reducing equivalents are involved, the bracketed and nonbracketed values are Gibbs free energies when the reducing equivalents are derived from either H2 or reduced NAD/NADP, respectively. For the reactions in which acetyl-CoA is synthesized from CH3-THF, HSCoA, and [CO], the Gibbs free energy is not known. Different values have been calculated for this reaction [e.g., 22 kJ mol 1 in Fuchs (1986) and 49 kJ mol 1 in Fuchs (1994)]. The value shown in the figure is an estimate that is based on the overall thermodynamic value of the pathway [i.e., 95 kJ mol 1 (Fuchs 1986, 1994)]. The parenthetically enclosed numbers (in blue) identify the different enzymes involved (see > Table 1.5 for characteristics of enzymes). The two enzymes responsible for the initial reduction of CO2 are formate dehydrogenase and acetyl-CoA synthase; reactions in which these two enzymes are involved are shaded in blue and green, respectively. Abbreviations: THF tetrahydrofolate, HSCoA coenzyme A, Pi inorganic phosphate, e reducing equivalent, and Co-Enzyme corrinoid enzyme

approximately one-third of the 14C from [U-14C]glucose is recovered in the exogenous CO2 pool when M. thermoacetica is grown on [U-14C]glucose; the other two-thirds of the 14C is recovered as [14C]acetate (D. R. Martin and H. L. Drake, unpublished data; see also Martin et al. 1985).

Exogenous CO2 can even be required for CO-dependent acetogenesis, a process that generates excess CO2 (Savage et al. 1987): 4CO þ 2H2 O ! CH3 COOH þ 2CO2

ð1:9Þ

27

28

1

Acetogenic Prokaryotes

a b

C6H12H6 (glucose) 2 ATP

2 CH3COCOOH (pyruvate)

2 NAD

2 CO2

Glycolysis

2 NADH (1)

2 E.TPP-CHOHCH3

E. TPP

(1)

2 HSCoA

8 Reducing Equivalents

2 Fdoi 2 Fdrod

2 CH3CO-SCoA

c

(2) 2 Pi

[6e–]

CO2

2 CH3COO-PO32− 2 ADP

Carbonyl Branch

Methyl Branch

ATP

[2e–]

CO2

(4–8) (11)

ADP + H2

(3) 2 ATP

THF

2 CH3COOH

[CO]

CH3-THF

Acetates I & II

(9–12)

HSCoA

CH3CO-SCoA (2) Pi CH3COO-PO32–

ADP ATP

(3) CH3COOH Acetate III

Acetyl-CoA Pathway

. Fig. 1.12 Transfer of reductant from glycolysis (Box A) and the oxidation of pyruvate (Box B) to the acetyl-CoA pathway (Box C). The overall scheme was elucidated from studies with Moorella thermoacetica. The parenthetically enclosed numbers (in blue) identify the different enzymes involved (see > Table 1.5 for characteristics of enzymes). Abbreviations: THF tetrahydrofolate, E.TPP enzyme.thiamine pyrophosphate, Fd ferredoxin, HSCoA coenzyme A, Pi inorganic phosphate, and e reducing equivalent

The acetogenic utilization of highly reduced one-carbon substrates is strictly dependent upon the availability of exogenous CO2. Thus, acetogenic cultures of M. thermoacetica and C. formicoaceticum cannot be maintained on methanol in the absence of supplemental CO2 (Hsu et al. 1990a; Matthies et al. 1993). The importance of CO2 to acetogens is exemplified by their ability to generate growth-essential CO2 equivalents from various compounds, including carboxylated lignin derivatives (e.g., vanillate; Hsu et al. 1990a, b). Acetogenesis is often referred to as a fermentation. Since the term ‘‘fermentation’’ implies that an organism uses a partially

oxidized carbonaceous intermediate as a terminal electron acceptor (e.g., when pyruvate is reduced to lactate in lactate fermentation), usage of this term for acetogenesis is not fully valid. For example, as outlined above, it appears that exogenous CO2 rather than that generated from the decarboxylation of pruvate is used as the terminal electron acceptor when glucose is metabolized. Likewise, the CO2 that is derived from the decarboxylation of aromatic compounds and subsequently used as a terminal electron acceptor in the acetyl-CoA pathway is fundamentally dissimilar to the use of an oxidized intermediate as a terminal electron acceptor in fermentation.

Acetogenic Prokaryotes

CO6H12O6 Glucose

Acetyl-CoA Pathway 2X 2 XH2

2 CH3COCOOH Pyruvate

½ CO3COOH + H2O CO2

Acetyl-CoA Pathway

2 H2O

2X

2 CO2

2 XH2

½ CO3COOH + H2O CO2

2 CH3COOH Acetate Sum: Glucose

3 Acetate

. Fig. 1.13 Importance of CO2 for the recycling of reduced electron carriers (XH2) during the acetogenic oxidation of glucose. Under certain growth conditions, exogenous CO2 is required for growth, even though CO2 is produced during catabolism (e.g., via the decarboxylation of pyruvate) (Modified from Drake 1994)

Furthermore, referring to acetogenesis as a fermentation does not properly reflect the chemiosmotic manner in which energy is conserved via the acetyl-CoA pathway and plasma membrane ATPases (see the section on > ‘‘Conservation of Energy and Bioenergetics’’ in this chapter).

Enzymology of the Acetyl-CoA Pathway The initial reactions on the methyl and carbonyl branches of the pathway are catalyzed by formate dehydrogenase and acetylCoA synthase, respectively. These two enzymes are responsible for reductive reactions that are thermodynamically very unfavorable (i.e., have positive Gibbs free energies under standard conditions; > Fig. 1.11). Formate dehydrogenase from M. thermoacetica is rich in metals (2 moles of selenium, 2 moles of tungsten, and approximately 36 moles of iron per mole of enzyme) and was the first enzyme in which tungsten was shown to be a biologically active trace metal (Yamamoto et al. 1983; Ljungdahl 1986). The tetrahydrofolate pathway facilitates the subsequent reduction of formate to the methyl level (Ljungdahl 1986). Acetyl-CoA synthase is a nickel-containing enzyme, reduces CO2 to the carbonyl (CO) level, and facilitates the synthesis of acetyl-CoA (Wood and Ljungdahl 1991). The acetyl-CoA that is formed by acetyl-CoA synthase is subsequently converted to acetate during catabolism or utilized in the synthesis of cell carbon during anabolism (> Fig. 1.11). The Gibbs free energy for the overall reduction of 2 moles of CO2 to 1 mole of acetate is approximately 95 kJ mol 1. The general properties of the enzymes involved in the acetyl-CoA pathway are outlined in > Table 1.5. Many of the enzymes from acetogens are extremely susceptible to inactivation by oxidation. For example, formate dehydrogenase and

1

acetyl-CoA synthase are among the most oxygen-sensitive enzymes known. Thus, the use of O2-free chambers has become routine for studying the enzymes central to acetogenesis. Although all of the enzymes of the acetyl-CoA pathway are important to the functionality of the pathway, several of the enzymes are worthy of special note: (a) Acetyl-CoA synthase catalyzes two reactions and is often referred to by two names (Diekert and Thauer 1978; Drake et al. 1980; Ragsdale et al. 1983; Wood and Ljungdahl 1991; Ragsdale 1994): CO dehydrogenase : COþH2 O ! CO2 þ 2Hþ þ 2e ð1:10Þ   Acetyl-CoA synthase : CH3 þ ½COŠ ð1:11Þ þ Coenzyme-A ! acetyl-CoA The discovery of the ability of this enzyme to catalyze reaction > 1.10 (Diekert and Thauer 1978) was paramount to later studies that resolved the physiological importance of the enzyme in reaction > 1.11 (see > Table 1.1 and the section on > ‘‘Historical Perspectives’’ in this chapter). In the acetyl-CoA pathway, acetyl-CoA synthase reduces CO2 to CO and subsequently fixes this CO2-derived carbon (i.e., CO) into an organic form (i.e., in acetyl-CoA). Acetyl-CoA synthase catalyzes the thermodynamically least favorable reaction in the pathway (> Fig. 1.11), a fact that might partially explain why this enzyme can represent up to 2 % of the soluble cell protein of an acetogen (Ragsdale et al. 1983). That acetyl-CoA synthase was discovered as CO dehydrogenase resulted in the acetyl-CoA pathway being sometimes referred to as the ‘‘CO dehydrogenase pathway.’’ This nomenclature is less than ideal as it does not accurately portray the physiological function of the enzyme in the pathway and also does not differentiate the enzyme (and thus the pathway) from the CO dehydrogenase used by aerobic carboxydotrophs that grow via the oxidation of CO to CO2 (Meyer 1988; Meyer et al. 1993, 2000). Recent studies on the crystal structure of the enzyme from M. thermoacetica have shown that the a subunits of acetyl-CoA synthase display both closed and open conformations, and that the active form of the enzyme has an Ni-Ni-[Fe4-S4] cluster at the active site (Darnault et al. 2003). Copper, once thought to be a part of this nickelcontaining metal cluster (Doukov et al. 2002), is an inhibitor of acetyl-CoA synthase and not a component of the catalytically active enzyme (Bramlett et al. 2003; Darnault et al. 2003). A nickel insertase is involved in the biosynthesis of acetyl-CoA synthase (Loke and Lindahl 2003). (b) Formate dehydrogenase (Yamamoto et al. 1983) and formyltetrahydrofolate synthetase (Lovell et al. 1990) are also centrally important because formate dehydrogenase ‘‘reductively fixes’’ CO2 to formate, and formyltetrahydrofolate synthetase subsequently ‘‘covalently fixes’’ this CO2-derived carbon (i.e., formate) into an organic form on the methyl branch of the pathway. A functional gene

29

30

1

Acetogenic Prokaryotes

. Table 1.5 Properties of the enzymes in the acetyl-CoA pathwaysa Number in the figuresa Enzyme

Mr

1

Pyruvate-Fd oxidoreductase

225,000 a2

Oxidation/decarboxylation of pyruvate to acetyl-CoA

19

Drake et al. (1981a)

2

Phosphotransacetylase

88,100 a4

Conversion of acetyl-CoA to acetylphosphate

1+9

Drake et al. (1981a)

3

Acetate kinase

60,000 n.r.

Conversion of acetyl-CoA and ADP to acetate and ATP

4

Formate dehydrogenase

340,000 a2b2 (96,000 NADPH-dependent reduction of CO2 1 + 3 (+22) and 76,000) to formate

5

Formyltetrahydrofolate (HCO-THF) synthetase

240,000 a4

Conversion of formate to HCO-THF

1

6, 7

Methenyltetrahydrofolate (CH-THF) cyclohydrolase and methylenetetrahydrofolate (CH2-THF) dehydrogenase complex

55,000 a2

HCO-THF converted to CH-THF, and CH-THF reduced to CH2-THF

1 4 ( 5)

8

Methylenetetrahydrofolate (CH2-THF) reductase

9

Methyltransferase

58,900 a2

Transfer of methyl unit from CH3-THF to coninoid enzyme

Drake et al. (1981a, b)

10

Coninoid enzyme

89,000 ab (34,000 and 55,000)

Methylation of acetyl-CoA synthase

Hu et al. (1984)

11

Acetyl-CoA synthase

12

Acetyl-CoA synthase disulfide reductase

Subunit composition Function in pathwaysa

289,000 a8

Reduction of CH2-THF to CH3-THF

Primary/ historical DGro (kJ mol 1)b reference

13

Schaupp and Ljungdahl (1974)

8

Mayer et al. (1982) 23

40 ( 22)

440,000 a3b3 (78,000 Reduction of CO2 to the CO level, and +20 (+38) and 71,000) synthesis of acetyl-CoA from CH322, 49d (81,730 and THF, CO, and CoASH 72,928)c 225,000 a4

Reduction of disulfide of CoA binding site of acetyl-CoA synthase

Yamamoto et al. (1983)

O’Brien et al. (1973)

Park et al. (1991)

Ragsdale et al. (1983) Diekert and Ritter (1983) Pezacka and Wood (1986)

Abbreviations: n.r. not reported, THF tetrahydrofolate, CoA coenzyme A, and Fd ferredoxin See > Figs. 1.11 and > 1.12. Enzymes indicated have been purified from M. thermoacetica b Gibbs free energies (Gor) have been rounded off and were obtained from Thauer et al. (1977) and Fuchs (1986). Parenthetical values for reactions catalyzed by oxidoreductases are the Gibbs free energies when the reducing equivalents are derived from reduced NAD/NADP rather than H2 c Based on amino acid composition. Current information indicates that the correct composition is a2b2 d Actual value is unknown. Different values have been calculated for this reaction [e.g., 22 kJ mol 1 in Fuchs (1986) and 49 kJ mol 1 in Fuchs (1994)] a

probe for the detection of acetogens is based on the gene sequence of formyltetrahydrofolate synthetase (Lovell and Hui 1991; Lovell 1994; Leaphart and Lovell 2001; see the section on > ‘‘Detection of Acetogens’’ in this chapter). (c) Under chemolithoautotrophic conditions (e.g., during growth on H2-CO2), the pathway must not only fix carbon but also conserve energy. Reactions that appear to be associated with this conservation of energy are facilitated by methlylenetetrahydrofolate reductase and methyltransferase (Clark and Ljungdahl 1984; Park et al. 1991; Wohlfarth and Diekert 1991; Mu¨ller et al. 2001; Mu¨ller 2003; see the section on > ‘‘Conservation of Energy and Bioenergetics’’ in this chapter).

(d) H2-dependent growth under chemolithotrophic conditions is considered to be a hallmark of acetogens, and this capability requires the activation of H2-derived reductant via hydrogenase. Though seldom highlighted in the pathway, hydrogenase thus catalyzes the first step in the chemolithoautotrophic fixation of CO2 (i.e., without utilizable reductant, CO2 cannot be fixed). Though the activities and properties of hydrogenases from different acetogens have been documented (e.g., Braun and Gottschalk 1981; Kellum and Drake 1984; Ragsdale and Ljungdahl 1984; Dobrindt and Blaut 1996; Drake et al. 1997), relatively little information has been published on hydrogenases from autotrophically grown acetogens. Acetogens can contain

1

Acetogenic Prokaryotes

multiple hydrogenases (Kellum and Drake 1984), and levels of hydrogenase activity in the membrane can increase when acetogens are cultivated at the expense of H2-CO2 (Braus-Stromeyer and Drake 1996), indicating that the function and intracellular localization of hydrogenases in acetogens are affected by cultivation conditions. (e) Carbonic anhydrase catalyzes the following reversible reaction (Lindskog et al. 1971; Karrasch et al. 1989; Albers and Ferry 1994; Kisker et al. 1996; Vandenberg et al. 1996): CO2 þ H2 O $ HCO3 þ Hþ

ð1:12Þ

This enzyme has been demonstrated in numerous acetogens, including A. woodii and S. silvacetica (Braus-Stromeyer et al. 1997). Carbonic anhydrase is widespread in nature, occurs in an extensive number of organisms, including humans, plants, and prokaryotes, and has multiple functions, including pH homeostasis, facilitated diffusion of CO2, interconversion of CO2 and HCO3 , and ion transport. Since CO2 is important to the acetylCoA pathway, one physiological function of carbonic anhydrase in acetogens might be to increase intracellular levels of CO2. Carbonic anhydrase has been purified approximately 300-fold from A. woodii; that specific activities of carbonic anhydrase in A. woodii are very high in both autotrophically and organotrophically grown cells indicates that this enzyme is physiologically important during acetogenesis (Braus-Stromeyer et al. 1997).

Conservation of Energy and Bioenergetics Acetogens can conserve energy by substrate-level phosphorylation and chemiosmotic mechanisms. Under certain growth conditions, both processes can be utilized simultaneously. However, when acetogens grow chemolithoautotrophically (e.g., on H2-CO2), energy can only be conserved via a chemiosmotic mechanism. Substrate-Level Phosphorylation. Under homoacetogenic conditions, certain hexoses are converted stoichiometrically to acetate, and a net of four ATP (per hexose metabolized) is formed by substrate-level phosphorylation (> Figs. 1.3 and > 1.12). A point that is often overlooked when comparing acetogens to other anaerobes is that acetogens conserve more energy by substrate-level phosphorylation than do the more classic fermenters (> Table 1.6). For example, twice as much energy is conserved via substrate-level phosphorylation during the homoacetogenic metabolism of glucose than during glucosedependent ethanol fermentation. It should be noted that when hexoses are metabolized, the net energy that is conserved via substrate-level phosphorylation is only indirectly linked to the acetyl-CoA pathway (see panels A and B in both > Figs. 1.3 and > 1.12). The ability of acetogens to conserve more energy via substrate-level phosphorylation than certain other anaerobes (i.e., on a per-mole-substrate-utilized basis) might make acetogens more competitive under certain in situ conditions.

. Table 1.6 Amount of ATP formed via substrate-level phosphorylation (ATPSLP) during glucose-dependent acetogenesis and fermentative processes Process

Stoichiometry of the process

ATPSLP

Acetogenesis

Glucose ! 3 acetate

4

Butyrate fermentation

Glucose ! butyrate + 2CO2 + 2 H2

3

Bifidum fermentation

Glucose ! 1.5 acetate + lactate 2.5

Ethanol fermentation

Glucose ! 2 ethanol + 2CO2

2

Homolactate fermentation

Glucose ! 2 lactate

2

Heterolactate fermentation

Glucose ! lactate + ethanol + 1 CO2

Membranous Electrochemical Gradients and ATPases. Although the above comments suggest that the ability to conserve energy via substrate-level phosphorylation is important to the bioenergetics of acetogens, the net production of ATP via substrate-level phosphorylation does not directly occur in the acetyl-CoA pathway. During the reductive synthesis of acetate from CO2, one ATP is consumed when formate is activated and one ATP is gained at the level of acetate kinase, thus yielding a break even relative to net ATP gain via substrate-level events (> Fig. 1.11). In addition, the reduction of CO2 to the carbonyl level on the carbonyl branch is thermodynamically unfavorable and requires energy (estimated at one-third ATP equivalent; Diekert 1992). Thus, in the absence of chemiosmotic processes, the acetyl-CoA pathway does not directly yield net utilizable energy via substrate-level phosphorylation. However, cell yields of acetogens are in excess of cell yields that can be explained from energy conserved via substrate-level phosphorylation. One mole of ATP yields approximately 10 g dry weight of microbial biomass, and cell yields of acetogens (e.g., C. aceticum and A. woodii) are 50–70 g cell dry weight per mole of glucose equivalent (Andreesen et al. 1973; Tschech and Pfennig 1984). Since a maximal cell yield of 40 g cell dry weight per mole of glucose is predicted from substrate-level phosphorylation (4 moles ATPSLP per mole of glucose), the higher than expected cell yields indicate that acetogens conserve energy by chemiosmotic processes. The capacity of acetogens to grow chemolithoautotrophically on H2-CO2 likewise indicates that acetogens are capable of conserving energy via electron transport phosphorylation or chemiosmotic mechanisms (since there is no net ATP gain via substrate-level phoshorylation in the acetyl-CoA pathway). Acetogens appear to use two different mechanisms for conserving energy by chemiosmotic processes. One process involves the generation of a membranous proton gradient and the synthesis of ATP by proton-dependent ATPase (Ljungdahl 1994; > Fig. 1.14a), and the other process involves the generation of a membranous sodium gradient and the synthesis of ATP by sodium-dependent ATPase (Mu¨ller and Gottschalk 1994;

31

32

1 a

Acetogenic Prokaryotes Proton-dependent ATPase Out H2 +

b

Sodium-dependent ATPase

In

Out MT

H2ase

CH3-THF

H

THF

Co-E H+

E T S

[H]

[e−]

Na+

Na+ ACS

CH3-Co-E

2 CO2

CH3-ACS

Acetate ADP + Pi H+

ADP + Pi Na+

ATPase

ATPase ATP

ATP membrane

repressed during nitrate dissimilation (see the section on of the Acetyl-CoA Pathway and Other Metabolic Abilities’’ in this chapter) is closely interfaced to proton translocation by membranous menaquinone during the transport of electrons (Das and Ljungdahl 2003). Examples of acetogens that have membranous electron transport systems and proton-dependent ATPases include M. thermoacetica, M. thermoautotrophica, and S. sphaeroides. (b) Some acetogens require sodium for growth, motility, and the optimal synthesis of acetate under certain conditions (Geerligs et al. 1989; Heise et al. 1989; Yang and Drake 1990; Mu¨ller and Bowien 1995). For example, T. kivui requires sodium for chemolithoautotrophic growth but not for organotrophic growth (Yang and Drake 1990). In contrast, M. thermoacetica does not require sodium for either chemolithoautotrophic or organotrophic growth (Yang and Drake 1990). Although the biochemical mechanism is not fully resolved, a membrane-associated complex that is interfaced to the methyl branch of the acetyl-CoA pathway appears to facilitate the translocation of sodium ions across the cell membrane (Mu¨ller and Gottschalk 1994; Mu¨ller et al. 2001; Mu¨ller 2003). The joint reaction catalyzed by methyltransferase and the corrinoid enzyme appears to be centrally important in the translocation of sodium ions, and the model proposed in > Fig. 1.14b illustrates how the generation of a sodium gradient is interfaced to the synthesis of ATP via sodium-dependent ATPase (Reidlinger and Mu¨ller 1994a; Spruth et al. 1995). Although this model is only hypothetical, numerous observations indicate that it has a sound theoretical basis. The reader is referred to Mu¨ller et al. (2001), (2004) and Mu¨ller (2003) for a more thorough treatment of this topic. Acetobacterium woodii and T. kivui are examples of acetogens that use sodium translocation and sodium-dependent ATPases for the conservation of energy. (c) Sodium-proton antiporters may facilitate specific changes in the type of electrochemical gradient that is used to conserve energy in acetogens (Terracciano et al. 1987; Yang and Drake 1990; Mu¨ller and Gottschalk 1994). For example, harmaline, an inhibitor of sodium-proton antiporters, uncouples the growth of T. kivui from the synthesis of acetate under chemolithoautotrophic conditions (i.e., the H2-dependent production of acetate continues in the absence of growth when harmaline is added to the growth medium; Yang and Drake 1990). Such observations suggest that sodium-proton antiporters are important to the ability of acetogens to conserve energy under certain conditions. > ‘‘Regulation

In

membrane

. Fig. 1.14 Proton- (a) and sodium-dependent (b) mechanisms for the chemiosmotic conservation of energy by acetogens. Panel A is based on the work of Ljungdahl and coworkers, and Panel B is based on the work of Mu¨ller and coworkers. Abbreviations: H2ase hydrogenase, ETS membranous electron transport system that is composed of various electron carriers (the system facilitates both the transport of electrons and translocation of protons), ATPase ATP synthase, MT methyltransferase, Co-E corrinoid enzyme, THF tetrahydrofolate, e reducing equivalent, and ACS acetyl-CoA synthase

Mu¨ller et al. 2001; > Fig. 1.14b). Although the precise biochemical mechanisms for these two processes are not known, several generalizations can be made as to how these processes occur: (a) Acetogens are rich in electron carriers (e.g., ferredoxins, rubredoxins, quinones, and cytochromes), and certain acetogens have membranous electron transport systems that can generate proton gradients across the membrane which can subsequently be used by proton-dependent ATP synthases (ATPases) to synthesize ATP (Das et al. 1989, 1997; Kamlage and Blaut 1993a; Kamlage et al. 1993b; Ljungdahl 1994; Das and Ljungdahl 2003). Likewise, certain oxidoreductases (e.g., hydrogenase) may be membraneassociated and also involved in generating proton gradients across the membrane (Ljungdahl 1994; Drake et al. 1997; > Fig. 1.14a). The large, negative Gibbs free energy for the methylenetetrahydrofolate-reductase-mediated step in the acetyl-CoA pathway (> Table 1.5) suggests that this enzyme is associated with energy conservation (Wohlfarth and Diekert 1991). This enzyme can be membrane bound (Hugenholtz et al. 1987) and might function in close association with the other catalysts responsible for the synthesis of acetyl-CoA that might also be loosely associated with the membrane [e.g., acetyl-CoA synthase activity occurs in the cell membrane of M. thermoautotrophica (Hugenholtz et al. 1987; Hugenholtz and Ljungdahl 1989) and T. kivui (Braus-Stromeyer et al. 1996; Drake et al. 1997)]. The membranous b-type cytochrome that is linked to the activity of methylenetetrahydrofolate reductase and

Additional Perspectives on the Bioenergetics of Acetogens. Pyrophosphate is a utilizable source of energy for certain anaerobes (Liu et al. 1982; Varma and Peck 1983). Moorella thermoacetica forms and subsequently consumes intracellular pyrophosphate during growth (Heinonen and Drake 1988). However, no direct correlation has been established between the intracellular turnover of pyrophosphate and cellular bioenergetics of acetogens.

Acetogenic Prokaryotes

Acetogens display different growth efficiencies with identical substrates. For example, the cell yields of T. kivui on H2 and glucose are approximately twofold higher than those of M. thermoacetica (Daniel et al. 1990). The biochemical explanations for such differences have not been resolved. However, the different mechanisms by which acetogens conserve energy may in part be responsible for such differences in growth efficiencies. With Sporomusa, different cytochromes are utilized for H2- and betaine-derived reductant (Kamlage and Blaut 1993a; Kamlage et al. 1993b); thus, the engagement of different electron transport systems for different substrates might account for some of the differences in growth efficiencies (i.e., the bioenergetics of growth) of acetogens.

Occurrence of the Acetyl-CoA Pathway in Nonacetogenic Microorganisms Metabolic schemes that bear close biochemical resemblance to the acetyl-CoA pathway of acetogens are utilized by nonacetogenic bacteria (e.g., sulfate-reducing bacteria) and members of the domain Archaea (e.g., methanogens) for either the assimilation of CO2 (i.e., carbon) into biomass or the oxidation of acetate (> Table 1.7). Thus, many of the biochemical and physiological features of the acetyl-CoA pathway are widely distributed in the prokaryotes. It must be remembered that these metabolic processes are not exactly the same, and that different metabolic cofactors and enzymes are involved. Nonetheless, the general features of these different forms of the acetyl-CoA pathway are very similar. Different species of the sulfate-reducing bacteria have been observed to have acetogenic capabilities (Klemps et al. 1985; Min and Zinder 1990; Madsen and Licht 1992; Tasaki et al. 1992, 1993; Kuever et al. 1993, 1999; Christiansen and Ahring 1996; Sanford et al. 1996); however, definitive, enzymological information on this metabolic

1

capability is scant. The acetyl-CoA pathway is the dominant biological mechanism for the anaerobic oxidation of acetate (Fuchs 1990).

Diverse Physiological Talents of Acetogens The standard redox potential of the CO2/acetate, CO2/methane, and sulfate/sulfide half-cell reactions approximates 290, 240, and 220 mV, respectively. Thus, under standard conditions, the reductive synthesis of acetate from CO2 is thermodynamically less favorable than methanogenesis or the reduction of sulfate to sulfide. Such thermodynamic limitations are often cited to explain why acetogens are physiologically less competitive for available reductant than methanogens and sulfate-reducing bacteria. However, as outlined in this section, acetogens have very diverse metabolic abilities that would theoretically increase their in situ competitiveness.

Diverse Electron Donors Diverse substrates can be oxidized and deliver reductant to the acetyl-CoA pathway and the reductive synthesis of acetate (> Table 1.8). Oxidizable substrates include CO, H2, carbohydrates, alcohols, carboxylic acids, dicarboxylic acids, aldehydes, substituent groups of aromatic compounds, and numerous other organic and halogenated substrates. By virtue of their ability to use a wide range of substrates, acetogens have numerous trophic links to other organisms under in situ conditions. Most acetogens have not been observed to degrade complex polymers, such as lignin or cellulose. However, little effort has gone into finding such acetogenic isolates. The recently described acetogen ‘‘B. formatexigens,’’ isolated from human feces, utilized amorphous (cabbage) cellulose and carboxymethylcellulose when isolated but lost this ability upon storage under frozen

. Table 1.7 Functions of acetyl-CoA synthase and the acetyl-CoA pathway in obligate anaerobesa

a

Group

Process

Acetyl-CoA forming

Acetogens

CO2 + [H] ! [acetyl-CoA] ! acetate + Drake (1994) biomass

Acetyl-CoA forming

Autotrophic methanogens, e.g., Methanobacterium thermoautotrophicum

CO2 + [H] ! [acetyl-CoA] ! ! biomass

Acetyl-CoA forming

Autotrophic S-reducers, e.g., Desulfobacterium autotrophicum

CO2 + [H] ! [acetyl-CoA] ! biomass Fuchs (1994)

Acetyl-CoA degrading

Acetate-oxidizing methanogens, e.g., Methanosarcina barkert

Acetate ! [acetyl-CoA] ! CO2 + CH4 Ferry (1994)

Acetyl-CoA degrading

Acetate-oxidizing S-reducers, e.g., Desulfotomaculum acetoxidans

Acetate ! [acetyl-CoA] ! CO2 + [H]

Fuchs (1994)

Bidirectional

Bidirectional acetogens, e.g., ‘‘Reversibacterium strain AOR’’

Acetate ⇄ [acetyl-CoA] ⇄ CO2 + [H]

Zinder (1994)

[H] is reductant. Acetyl-CoA is bracketed to indicate that it is an intracellular intermediate See also Fuchs (1986, 1989) and Wood and Ljungdahl (1991) Modified from Drake (1992, 1994) b

Referencesb

Function

Whitman (1994)

33

34

1

Acetogenic Prokaryotes

. Table 1.8 Representative growth-supportive substrates and overall substrate/product stoichiometries of acetogens that can form acetate as their predominant reduced producta Substrate

Overall stoichiometry for acetate production

Representative acetogen

Acetoin

2 CH3COCHOHCH3 + 2 CO2 + 2 H2O ! 5 CH3COOH

Acetobacterium carbinolicum

Alcoxyethanols

4 RO-CH2CH2OH + 2 CO2 2 H2O ! 4 ROH + 5 CH3COOH

Acetobacterium malicum

e.g., 2-Methoxyethanol

4 CH3OCH2CH2OH + 2 CO2 + 2 H2O ! 4 CH3OH + 5 CH3COOH

Acetobacterium malicum

e.g., 2-Ethoxyethanol

4 C2H5OCH2CH2OH + 2 CO2 + 2 H2O ! 4 C2H5OH + 5 CH3COOH

Acetobacterium malicum

2,3-Butanediol

4 CH3CHOHCHOHCH3 + 6 CO2 + 2 H2O ! 11 CH3COOH

Clostridium magnum

Cellobiose

C12H22O11 + H2O ! 6 CH3COOH

Ruminococcus productus

Citrate

4 C6H8O7 6+ 2 H2O ! 9 CH3COOH + 6 CO2

Clostridium magnum

CO

4 CO + 2 H2O ! CH3COOH + 2 CO2

Moorella thermoacetica Clostridium formicoaceticum

Ethanol

2 CH3CH2OH + 2 CO2 ! 3 CH3COOH

Formate

4 HCOOH ! CH3COOH + 2 CO2 + 2 H2O

Moorella thermoacetica

Fructose

C6H12O6 ! 3 CH3COOH

Clostridium formicoaceticum

Glucose

C6H12O6 ! 3 CH3COOH

Clostridium thermoaceticum

Glycerol

4 HOCH2CHOHCH2OH + 2 CO2 ! 7 CH3COOH + 2 H2O

Acetobacterium carbinolicum

Glycolate

4 CH2OHCOOH ! 3 CH3COOH + 2 CO2 + 2 H2O

Moorella thermoacetica

Glyoxylate

2 HOCCOOH ! CH3COOH + 2 CO2

Moorella thermoacetica

H2 + CO2

4 H2 + 2 CO2 ! CH3COOH + 2 H2O

Clostridium aceticum

H2 + CO

2 H2 + 2 CO ! CH3COOH

Moorella thermoacetica

H2 + formate

2 H2 + 2 HCOOH ! CH3COOH + 2 H2O

Moorella thermoacetica

4-Hydroxybenzaldehyde 4, 4-Hydroxybenzaldehyde + 2 CO2 + 2 H2O ! CH3COOH + 4-hydroxybenzoate Clostridium formicoaceticum Malate

2 HOOCCHOHCH2COOH ! 3 CH3COOH + 2 CO2

Methanol

4 CH3OH + 2 CO2 ! 3 CH3COOH + 2 H2O

Clostridium magnum Moorella thermoacetica

Methoxyacetate

4 CH3OCH2COOH + 2 CO2 + 2 H2O ! 3 CH3COOH + 4 HOCH2COOH

Acetobacterium sp. RMMac1

Methoxylated aromatics 4 Aromatic-[OCH3] + 2 CO2 + 2 H2O ! 4 aromatic-[OH] + 3 CH3COOH

(Many acetogens)

e.g., Syringate

2 Syringate[-OCH3]2 + 2 CO2 + 2 H2O ! 2 gallate[-OH]2 + 3 CH3COOH

Acetobacterium woodii

e.g., Vanillate

4 Vanillate[-OCH3] + 2 CO2 + 2 H2O ! 4 protocatechuate[-OH] + 3 CH3COOH

Acetobacterium woodii

e.g., Syringate + H2-CO2 Syringate[-OCH3]2 + 2 CO2 + 2 H2 ! gallate[-OH]2 + 2 CH3COOH

Strain SS1

Methyl chloride

4 CH3Cl + 2 CO2 + 2 H2O ! 3 CH3COOH + 4 HCl

‘‘Acetobacterium dehalogenans’’

Oxalate

4 HOOCCOOH ! CH3COOH + 6 CO2 + 2 H2O

Moorella thermoacetica

Pyruvate

4 CH3COCOOH + 2 H2O ! 5 CH3COOH + 2 CO2

Moorella thermoacetica

Xylose

2 C5H10O5 ! 5 CH3COOH

Moorella thermoacetica

a

No distinction is made (a) between acids and their dissociated salt forms or (b) between CO2 and its carbonate or bicarbonate forms. Although only one representative acetogen is listed per substrate, many acetogens may be capable of utilizing each substrate (e.g., almost all acetogens can synthesize acetate from H2/CO2 or methoxylated aromatic compounds)

conditions (Wolin and Miller 1994; Wolin et al. 2003). Another acetogen phylogenetically closely related to M. thermoacetica, strain F21 (isolated in a roll tube containing Avicel® [crystalline cellulose]), is cellulolytic and has carboxymethylcellulase and xylanase activities (Karita et al. 2003). The cellulolytic capabilities of these two organisms suggest that certain acetogens may be able to degrade certain polymers. An organism referred to as ‘‘M. thermoautotrophica’’ (C. thermoautotrophicum at the time of publication) was reported to degrade inulin, a large (Mr ca. 5,000; Budavari 1989) storage polysaccharide of plants (Drent and Gottschal 1991).

However, the product profile (1.0 hexose ! 0.4 formate + 0.7 acetate + 1.3 ethanol + 1.0 H2 + 1.0 CO2 + 0.6 cell carbon; Drent and Gottschal 1991) of this isolate is very inconsistent with that of M. thermoautotrophica (Wiegel et al. 1981) and also of acetogens. In the absence of additional information, it cannot be assumed that this organism is either M. thermoautotrophica or an acetogen. The ability to utilize methoxyl groups of aromatic compounds (e.g., vanillate) is a widespread metabolic potential of acetogens (Bache and Pfennig 1981; Frazer and Young 1985; Daniel et al. 1991; Frazer 1994). > Figure 1.15 illustrates how

Acetogenic Prokaryotes

Terminal Portion of Methyl Branch 3X

Reverse Methyl Branch

Carbonyl Branch

3 CH3-THF

X

CH3-THF

3 CO2

Demethylation 3 X-CH3

1

Demethylation THF

3 THF 6 e−

X-CH3

ATP COOH

3 [CO]

3 CH3-[Co-Protein]

CO2 3 Acetyl-CoA

3 Acetylphosphate 3 ATP

X = chloride or aromatic compound (corrinoid-bound methyl unit might be an intermediate in the demethylation step)

3 Acetate

. Fig. 1.15 Pathway illustrating how methyl groups from methoxylated aromatic compounds (e.g., vanillate) or methyl chloride can be metabolized by acetogens. Abbreviations: THF tetrahydrofolate, CoA coenzyme A, e , reducing equivalent, and Co-Protein corrinoid enzyme

methyl groups can be metabolized by acetogens. This metabolic scheme is based on the work of several laboratories (see below), and it must be remembered that the demethylase system varies, depending on the substrate that is demethylated. Furthermore, there may be secondary ATP-dependent activation steps that maintain corrinoid proteins (which may be involved in demethylation and transfer of methyl groups) in a reduced, active form (Kaufmann et al. 1997, 1998). The methylotrophic potential of acetogens to utilize methyl groups makes physiological sense, given the importance of methyl-level intermediates in the acetyl-CoA pathway. An unusual and apparently highly specialized methylotrophic metabolism is seen in the ability of ‘‘A. dehalogenans’’ to utilize methyl chloride according to the following reaction (Traunecker et al. 1991): 4CH3 Cl þ 2CO2 þ 2H2 O ! 3CH3 COOH þ 4HCl

ð1:13Þ

Methyl chloride is dehalogenated by methyl chloride dehalogenase, which yields chloride and methyltetrahydrofolate (Meßmer et al. 1993, 1996). The methyl unit of methyltetrahydrofolate is oxidized to CO2 by reversal of the methyl branch of the acetyl-CoA pathway, the reductant being subsequently used for the reduction of CO2 to CO on the carbonyl branch of the pathway (> Fig. 1.15). ‘‘Acetobacterium dehalogenans’’ also utilizes methoxylated aromatic compounds, and the O-demethylase of ‘‘A. dehalogenans’’ consists of two distinct methyltransferases. Methyltransferase I O-demethylates the methoxylated aromatic compound and transfers the methyl group to a corrinoid protein. Methyltransferase II transfers the methyl group of the methylated corrinoid protein

to tetrahydrofolate (Kaufmann et al. 1997, 1998). The methyltetrahydrofolate is metabolized in the same manner as the methyltetrahydrofolate that is derived from methyl chloride, i.e., the methyl group of methyltetrahydrofolate (1) serves as a methyl donor at the terminal stage of the methyl branch of the acetyl-CoA pathway or (2) is oxidized, and thus serves as a source of reductant for the reductive formation of CO on the carbonyl branch of the acetyl-CoA pathway (> Fig. 1.15). Acetobacterium woodii and M. thermoacetica appear to utilize O-demethylating systems that are similar to that of ‘‘A. dehalogenans’’ (Berman and Frazer 1992; Frazer 1994; Naidu and Ragsdale 2001). An O-demethylating methyltransferase system that is involved in the metabolism of methoxyl groups of aromatic compounds has also been characterized from H. foetida (Kreft and Schink 1997). Holophaga foetida is an unusual acetogen that can (1) degrade aromatic rings (Bak et al. 1992; Kreft and Schink 1993; Liesack et al. 1994) and (2) methylate sulfide to dimethylsulfide via a non-energy-conserving, methyltransferase-mediated reaction (Kappler et al. 1997). The O-demethylating methyltransferase system of H. foetida does not appear to be identical to the methyltransferase system characterized from ‘‘A. dehalogenans’’ (Kaufmann et al. 1998). O-demethylating methyltransferases of acetogens are inducible and may be either specific or nonspecific for the methoxyl group that is O-demethylated (Wu et al. 1988; Daniel et al. 1991; Ha¨ggblom et al. 1993; Kreft and Schink 1997). Certain acetogens can O-demethylate halogenated aromatic compounds; however, the residual aromatic compound may not be subject to dehalogenation (Ha¨ggblom et al. 1993).

35

36

1

Acetogenic Prokaryotes

. Table 1.9 Terminal electron acceptors used by acetogensa Electron acceptor

Reduced end product

Representative acetogen

References Misoph and Drake (1996)

Acetaldehyde

Ethanol

Ruminococcus productus

Carbon dioxide

Acetate

All acetogens

Wood and Ljungdahl (1991)

Dimethylsulfoxide

Dimethylsulfide

Moorella thermoacetica

Beaty and Ljungdahl (1991)

Fumarate

Succinate

Clostridium aceticum

Matthies et al. (1993)

Methoxylated phenylacrylates

Methoxylated phenylpropionates

Acetobacterium woodii

Bache and Pfennig (1981)b

Nitrate

Nitrite

Moorella thermoacetica

Seifritz et al. (1993)c

Nitrite

Ammonium

Moorella thermoacetica

Seifritz et al. (2003)

Protons

Molecular hydrogen

Acetobacterium woodii

Winter and Wolfe (1980)d

Pyruvate

Lactate

Ruminococcus productus

Misoph and Drake (1996)

Thiosulfate

Sulfide

Moorella thermoautotrophica

Beaty and Ljungdahl (1990)

a

No known acetogen is able to use all of the electron acceptors listed. The ability to use a particular electron acceptor is conditional and is dependent upon in situ conditions and the acetogen in question b See also Tschech and Pfennig (1984), Hansen et al. (1988), and Imkamp and Mu¨ller (2002) c See also Fro¨stl et al. (1996) and Seifritz et al. (2002) d The H2 formed can be used by an H2-utilizing methanogen for interspecies hydrogen transfer

The acetyl-CoA pathway can be considered a one-carbon pathway, in that each of the two branches of the pathway facilitates the reduction of one-carbon substrates. Thus, entry-level, onecarbon substrates are particularly well suited for use by acetogens, especially under cosubstrate conditions. For example, CO and the methoxyl group of aromatic compounds can be simultaneously and readily metabolized via the carbonyl or methyl branches of the acetyl-CoA pathway, respectively. Methyl groups of osmolytes can also be metabolized by acetogens (Jansen and Hansen 2001). Certain acetogens also utilize two-carbon substrates, such as oxalate, glyoxylate, and glycolate; however, how these twocarbon molecules are metabolized is not well resolved (Daniel and Drake 1993; Daniel et al. 2004; Seifritz et al. 1999, 2002). Acetogens are specialized in the use of short-chain substituent groups of aromatic compounds, and the use of such substituent groups can have regulatory effects on the flow of carbon and reductant. For example, use of the aldehyde group of certain lignin-derived aromatic compounds by C. formicoaceticum inhibits the cell’s ability to use fructose (i.e., in the presence of both fructose and 4-hydroxybenzaldehyde, reductant for the acetyl-CoA pathway and the reduction of CO2 are preferentially derived from the aldehyde group of 4-hydroxybenzaldehyde; Go¨ßner et al. 1994; Frank et al. 1998; Drake and Ku¨sel 2003, 2005).

Use of Diverse Terminal Electron Acceptors The acetyl-CoA pathway is the hallmark of acetogens, and, as outlined in the previous section > ‘‘Diverse Electron Donors,’’ diverse substrates can be oxidized by acetogens and thus utilized for the reductive synthesis of acetate from CO2. However, the acetyl-CoA pathway is not the only terminal-electron-accepting process utilized by acetogens (> Table 1.9). Thus, the trophic relationships and adaptation strategies of acetogens under in

situ conditions may not be solely determined by their acetogenic capabilities (i.e., their use of the acetyl-CoA pathway). A few generalizations can be made about the ability of acetogens to engage in these alternative metabolic processes: (a) Almost all known acetogens can use more than one terminal electron acceptor and, therefore, can produce other reduced end products in addition to acetate (> Table 1.9). The capacity to utilize a particular terminal electron acceptor is dependent on the availability of both reductant and terminal electron acceptor. Thus, homoacetogenesis is usually a conditional capacity of an acetogen, and referring to an acetogen as a homoacetogen is almost always a misnomer. (b) The availability of CO2 can determine how a particular substrate is metabolized by an acetogen. Indeed, for acetogenic growth, exogenous (i.e., supplemental) CO2 is often essential for growth (see the section on > ‘‘CO2 as Terminal Electron Acceptor and the Concept of Fermentation’’ in this chapter). However, even when exogenous CO2 is readily available, an acetogen may utilize other terminal electron acceptors, either in preference to or simultaneously with CO2. For example, R. productus simultaneously reduces CO2 to acetate and phenylacrylates to phenylpropionates (Misoph et al. 1996b), and also forms lactate as a fermentation end product concomitant to the reductive synthesis of acetate from CO2 during the metabolism of fructose (Misoph and Drake 1996a). (c) H2-dependent acetogenesis is thermodynamically difficult because of the thermodynamic constraints of the entry-level redox reactions for CO2 in the acetyl-CoA pathway (> Fig. 1.11). For example, the standard redox potential of the CO2/CO half-cell ( 520 mV) is approximately 100 mV more negative than that of the 2 H+ + 2e /H2 half-cell ( 420 mV). However, the capacity of an acetogen to consume H2 can be significantly improved when an alternative

Acetogenic Prokaryotes

(d)

(e)

(f)

(g)

electron acceptor is utilized. The H2 threshold of A. woodii is 520 parts per million (ppm) when CO2 is utilized as the terminal electron acceptor but decreases to 3 ppm when aromatic acrylates (e.g., caffeate) are utilized as terminal acceptors (Cord-Ruwisch et al. 1988). Likewise, the amount of biomass synthesized per mole of H2 consumed increases eightfold when nitrate instead of CO2 is utilized as the terminal electron acceptor by M. thermoautotrophicum during growth on low concentrations of H2 (Fro¨stl et al. 1996). Thus, the capacity of an acetogen to compete for H2 can increase significantly when alternative electron acceptors are utilized. The use of diverse electron acceptors by acetogens indicates that acetogens can accommodate a wide range of redox conditions. For example, nitrate is the preferred terminal electron acceptor of the classic acetogen M. thermoacetica (Seifritz et al. 1993, 2002; Fro¨stl et al. 1996), suggesting that this acetogen does not require stringently reduced conditions (i.e., the standard redox potential of the nitrate/ ammonium half-cell approximates +360 mV while that of the CO2/acetate half-cell approximates 290 mV). Indeed, M. thermoacetica is easily isolated from aerated soils that have fluctuating redox conditions (Go¨ßner and Drake 1997; Go¨ßner et al. 1998, 1999). The use of an alternative electron acceptor can have regulatory effects on the acetyl-CoA pathway (see the section on > ‘‘Regulation of the Acetyl-CoA Pathway and Other Metabolic Abilities’’ in this chapter). The use of an alternative electron acceptor can conserve energy and increase the general efficiency of growth. For example, when A. woodii is grown at the expense of methanol, growth yields are significantly greater when aromatic acrylates (e.g., caffeate or ferulate) are used as terminal electron acceptors rather than CO2 (Tschech and Pfennig 1984). The sodium-dependent reduction of aromatic acrylates by A. woodii is coupled to the synthesis of ATP (Hansen et al. 1988; Imkamp and Mu¨ller 2002). Likewise, growth yields of M. thermoacetica and M. thermoautotrophica increase significantly when reductant flow is directed to the dissimilation of nitrate rather than the reduction of CO2 to acetate (Seifritz et al. 1993; Fro¨stl et al. 1996). In fact, ethanol and propanol are not acetogenic substrates for these two acetogens, yet both alcohols are oxidized and growth supportive when nitrate is dissimilated (Fro¨stl et al. 1996). Thus, the use of alternative electron acceptors by an acetogen can increase the likelihood that certain compounds can be oxidized and be growth supportive. Energy might not always be conserved by the reduction of a given terminal electron acceptor. For example, H2 is produced as an end product by M. thermoacetica even though the production of H2 is very likely not directly linked to the conservation of energy (Martin et al. 1983, 1985; Fro¨stl et al. 1996). However, the ability of certain acetogens (e.g., A. woodii) to produce H2 as a substrate for the interspecies transfer of H2 indicates that the production of H2 can be coupled to the conservation of energy under certain in situ

1

conditions (Winter and Wolfe 1980; Cord-Ruwisch and Ollivier 1986; Heijthuijsen and Hansen 1986). (h) Certain acetogens have the ability to reductively dehalogenate low-molecular-weight halogenated compounds (e.g., carbon tetrachloride and tetrachloroethylene; Egli et al. 1988; Freedman and Gossett 1991; Traunecker et al. 1991; Holliger and Schraa 1994; Terzenbach and Blaut 1994; Hashsham and Freedman 1999). Thus, halogenated compounds can be an electron sink for acetogens. Reductive dehalogenation occurs concomitantly with acetogenesis. However, results to date indicate that reductive dehalogenation (1) is not directly linked to an enzymatic process, (2) is due to chemical reactions with reduced corrinoids that normally serve as cofactors during normal acetogenic metabolism, and (3) does not conserve energy.

Regulation of the Acetyl-CoA Pathway and Other Metabolic Abilities Acetogenesis was initially thought of as a constitutive trait of the classic acetogen M. thermoacetica. Since our understanding of acetogens was largely influenced by the decades of work on this acetogen (see the section on > ‘‘Historical Perspectives’’ in this chapter), it has only been in more recent times that we have learned that this view is a misconception regarding not only M. thermoacetica but acetogens in general. Although the molecular details are not yet well understood, it is now clear that many of the diverse metabolic processes of acetogens are regulated. Indeed, because acetogens have so many diverse metabolic capabilities, it is essential that these capabilities can be regulated. Numerous examples with many acetogens can be cited to reinforce this fact: (1) hydrogenase, formate dehydrogenase, and acetyl-CoA synthase activities are significantly influenced by growth substrates (Braun and Gottschalk 1981; Kellum and Drake 1986; Daniel et al. 1990; Lux and Drake 1992), (2) the capacity to utilize the carboxyl, methoxyl, and acrylate groups of certain aromatic compounds is inducible (Wu et al. 1988; DeWeerd et al. 1988; Hsu et al. 1990b; Imkamp and Mu¨ller 2002), (3) the ability to dehalogenate methyl chloride is inducible (Meßmer et al. 1993, 1996), and (4) electron transport systems are regulated (Kamlage et al. 1993b; Fro¨stl et al. 1996; Seifritz et al. 2002, 2003). Several genes encoding enzymes of the acetyl-CoA pathway of M. thermoacetica have been cloned and successfully expressed in Escherichia coli (Morton et al. 1992); however, regulation is not well understood at the molecular level. Relatively few studies have directly assessed the regulation of the acetyl-CoA pathway. The first evidence that the acetyl-CoA pathway is subject to regulation was obtained with A. woodii. Acrylate groups of certain aromatic compounds (e.g., caffeate) can be used as an alternative electron acceptor when A. woodii (grown at the expense of methanol) results in the total shutdown of the cell’s ability to produce acetate (Tschech and Pfennig 1984). Thus, the use of an alternative electron acceptor can have regulatory effects on the acetyl-CoA pathway.

37

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Acetogenic Prokaryotes

Methyl Branch

Carbonyl Branch

CO2

CO2 4 e−

[CH2]-THF Membranous Cytochrome b

2 e−

?

2 e−

Nitrate Block [CH3]-THF Site 1

Nitrate Block Site 2

(THF) OCH3

Methyl By-Pass

Carbonyl By-Pass

Co

Tolerance to Oxic Conditions and Metabolism of O2

OH [Co]

[CH3]-[Co-Protein] Acetyl-CoA Synthase

Carbon flows primarily towards acetate when nitrate is not present.

CH3COOH

level of acetyl-CoA synthase on the carbonyl branch of the pathway by an unknown mechanism that might also involve the flow of reductant to CO2 (Nitrate Block Site 2; > Fig. 1.16). Since acetyl-CoA synthase and hydrogenase can be associated with the cell membrane of certain acetogens (Hugenholtz et al. 1987; Hugenholtz and Ljungdahl 1989; Braus-Stromeyer et al. 1996; Dobrindt and Blaut 1996; Drake et al. 1997), it could be that the membranous b-type cytochrome and/or membranous hydrogenase is important for electron flow on the carbonyl branch of the pathway. Although the different functions of hydrogenase(s) during growth of acetogens on sugars are unknown, this oxidoreductase might be essential for the proper flow of sugar-derived reductant during the reduction of CO2 to acetate (Drake and Ku¨sel 2003).

Acetyl-CoA

When nitrate is present, carbon flows towards cell carbon when nitrate block is by-passed with methoxyl groups of aromatic compounds and CO.

Cell Carbon

. Fig. 1.16 Pathway illustrating where nitrate blocks the flow of carbon in the acetyl-CoA pathway when nitrate is dissimilated to ammonium by Moorella thermoacetica. Abbreviations: THF tetrahydrofolate, CoA coenzyme-A, and Co-Protein corrinoid enzyme. The dissimilation of nitrite to ammonium appears to have the same effect (Seifritz et al. 2002a) (Modified from Drake et al. 2003)

How the acetyl-CoA pathway can be regulated is best understood from information obtained with M. thermoacetica. The dissimilation of nitrate represses the function or engagement of the acetyl-CoA pathway by repressing the synthesis of a membranous b-type cytochrome that is essential for the formation of methyltetrahydrofolate on the methyl branch of the pathway (Nitrate Block Site 1; Fro¨stl et al. 1996; Drake et al. 2002; > Fig. 1.16). The amount of certain enzymes of the acetylCoA pathway might be less under nitrate-dissimilating conditions than under acetogenic conditions (Arendsen et al. 1999). However, all of the enzymes responsible for the flow of carbon in the acetyl-CoA pathway are expressed and functionally present in the cell when the cell is dissimilating nitrate but not reductively synthesizing acetate from CO2 (Fro¨stl et al. 1996). Thus, the membranous b-type cytochrome is a key element in regulating the flow of both reductant and carbon in the acetyl-CoA pathway. Hydrogenase activity is strongly repressed when cells dissimilate nitrate, and a second metabolic block occurs at the

Acetogens have been mostly isolated from habitats that are anoxic (> Table 1.2) and are routinely cultivated under anoxic conditions. Furthermore, many enzymes of the acetyl-CoA pathway are extremely sensitive to O2. Thus, acetogens have been classically referred to as strict anaerobes. However, acetogens are also present in aerated soils and colonize habitats with fluctuating redox conditions (e.g., the rhizosphere of sea grass). It should therefore come as no surprise that acetogens, like other so-called strict anaerobes [e.g., sulfate-reducing bacteria (Marschall et al. 1993; Johnson et al. 1997; Teske et al. 1998; Cypionka 2000)], are able to cope with oxidative stress. Indeed, acetogens isolated from aerated soils and the rhizosphere of sea grass have the ability to tolerate and consume O2 (Ku¨sel et al. 2001; Karnholz et al. 2002). Acetogens isolated from the termite gut can also tolerate transient, moderately oxic conditions (Boga and Brune 2003; Graber and Breznak 2004a). Although acetogens such as A. woodii, C. magnum, C. glycolicum RD-1, M. thermoacetica, and S. silvacetica can tolerate 0.5–6 % O2 (amount is dependent upon the acetogen and the conditions of incubation) in the headspace of culture tubes, the ability of acetogens to metabolize O2 does not appear to be directly coupled to the conservation of energy. The capacity of acetogens to metabolize O2 is likely used as a means to remove trace amounts of O2 (or toxic products derived from O2 [e.g., superoxides and peroxides]) from acetogen-colonized microniches that are subject to transient oxic conditions. Information to date suggests that there are three basic ways that acetogens can cope with oxic conditions: 1. When certain acetogens are challenged with oxic conditions, they make use of alternative electron acceptors and metabolically bypass the need to use the acetyl-CoA pathway. For example, the ability of M. thermoacetica to dissimilate nitrate negates the need of the cell to use CO2 as a terminal electron acceptor when conditions become more oxic. Another example is C. glycolicum RD-1, an aerotolerant acetogen that was isolated from sea grass roots and can tolerate up

Acetogenic Prokaryotes

Sugars Glycolysis, Decarboxylation, & Oxidation

Acetate + CO2

[Reductant]

Anoxic Conditions

CO2, Acetaldehyde

Acetogenesis & Fermentation

Acetate, Ethanol

Oxic Conditions

Acetaldehyde, Pyruvate, H+

Fermentation

Ethanol, Lactate, H2

. Fig. 1.17 Flow of sugar-derived reductant during the metabolism of Clostridium glycolicum RD-1. Under anoxic conditions, intracellular CO2 (taken up from exogenous sources or produced from the decarboxylation of pyruvate) and acetaldehyde (produced during catabolism) serve as terminal electron acceptors and are reduced to acetate and ethanol, respectively. Under oxic conditions, intracellular acetaldehyde, pyruvate, and protons (all produced during catabolism) serve as terminal electron acceptors and are reduced to ethanol, lactate, and H2, respectively (Modified from Ku¨sel et al. 2001)

to 4 % O2 in the headspace of shaken culture tubes (Ku¨sel et al. 2001). Clostridium glycolicum RD-1 is unusual in that it simultaneously utilizes acetogenesis and ethanol fermentation under anoxic conditions (> Fig. 1.17). However, when conditions become more oxic, acetaldehyde, pyruvate, and protons are exclusively used as terminal electron acceptors, and ethanol, lactate, and H2 become the reduced end products (> Fig. 1.17). Thus, the ability of certain acetogens to cope with oxidative stress appears to be maximized when sugars or other fermentable substrates are available. Such findings indicate that certain acetogens can shift the flow of reductant toward catabolic processes that are less sensitive to O2 when conditions become more oxic. 2. Several acetogens have been examined at either the enzymeactivity level or gene level for the presence of enzymes that might be involved in the removal (i.e., metabolism) of O2 or its toxic by-products (> Table 1.10; it should be noted that, thus far, not very many acetogens have been examined for these enzymes). The enzymes that have been detected at either the activity or gene level include NADHoxidase, peroxidase, superoxide dismutase, rubredoxin

1

oxidoreductase, and rubrerythrin. Unlike aerobes that utilize superoxide dismutase, an enzyme that forms O2 (> Table 1.10), only one acetogen (C. glycolicum RD1, an acetogen that displays a very high tolerance to O2 [above]) has been found to have this enzyme (Ku¨sel et al. 2001). Clostridium glycolicum RD1 also contains peroxidase and NADH-oxidase. To date, catalase, which also forms O2, has only been detected in the termite isolates of A. longum, S. termitida, and S. aerivorans (Kane and Breznak 1991a; Boga and Brune 2003; Boga et al. 2003). Clostridium magnum, M. thermoacetica, and S. silvacetica contain peroxidase and NADH-oxidase activities but lack catalase and superoxide dismutase (Karnholz et al. 2002). Rubredoxin oxidoreductase and rubrerythrin are oxidative stress enzymes in certain sulfate-reducing bacteria (Lumppio et al. 2001; Kurtz 2003), and genes for similar proteins occur in M. thermoacetica (Das and Ljungdahl 2001). A flavoprotein/rubredoxin combination (derived from M. thermoacetica) functions as an NADH:O2 oxidoreductase (Silaghi-Dumitrescu et al. 2003). Although these enzymatic capabilities vary from one acetogen to another, it can be concluded that acetogens have the capacity to reductively remove O2, peroxide, and superoxide. 3. Close trophic relationships between acetogens and microaerophiles that consume trace amounts of oxygen are also a means by which acetogens might cope with transient oxic conditions (Go¨ßner et al. 1999). (This trophic relationship is discussed in the section on > ‘‘Metabolic Interactions in Pure Culture and Complex Ecosystems’’ in this chapter.)

Ecology of Acetogens There is a general paradox regarding the ecology of acetogens: although theoretical considerations suggest that acetogenesis should not be a highly competitive microbial process, acetogens occur in highly diverse habitats, and their activity can sometimes compete with and even overshadow that of other anaerobes. This section will evaluate both sides of this paradox and will focus on the metabolic interactions of acetogens in certain ecosystems.

Metabolic Interactions of Acetogens in Pure Cultures and Complex Ecosystems In anoxic environments, acetogens compete with primary fermenters for monomeric compounds that are derived from the degradation of polymers and with secondary fermenters for typical fermentation products such as lactate, ethanol, or H2 (McInerney and Bryant 1981). Acetogenesis yields more ATP per mole of sugar than classic fermentations do, which might increase the competitiveness of acetogens under certain conditions (see the section on > ‘‘Conservation of Energy and Bioenergetics’’ and > Table 1.6 in this chapter). However, in pure culture, acetogens tend to grow slower on sugars than do

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. Table 1.10 Possible oxidative stress enzymes in acetogens Enzyme

Reaction +

+

Detected

References Ku¨sel et al. (2001)

NADH-oxidase

O2 + 2 NADH + 2 H ! 2 NAD + 2 H2O

Activity

Peroxidase

H2O2 + XH2 ! X + 2 H2O

Activity

Superoxide dismutase

2 O2 + 2 H+ ! O2 + H2O2

Activity

Catalase

2 H2O2 ! O2 + 2 H2O

Activity

Karnholz et al. (2002) Ku¨sel et al. (2001) Karnholz et al. (2002) Ku¨sel et al. (2001) Kane and Breznak (1991) Boga et al. (2003) FprA/Hrba

O2 + 2 NADH + 2 H+ ! 2 NAD+ + 2 H2O +

Activity

Silaghi-Dumitrescu et al. (2003)

Rubredoxin oxidoreductase

e + O2 + 2 H ! H2O2

Gene

Das et al. (2001)

Rubrerythrin

H2O2 + XH2 ! X + 2 H2O

Gene

Das et al. (2001)

Abbreviations: NAD nicotinamide adenine dinucleotide, NADH reduced nicotinamide adenine dinucleotide, FprA type A flavoprotein; and Hrb, rubredoxin a This combination also displays NO reductase activity

classic fermenters (e.g., Clostridium butyricum; Schink 1994). Nonetheless, acetogenesis is the most important anaerobic glucose-consuming process in anoxic paddy soils and lake sediments (Krumbo¨ck and Conrad 1991). When H2 is utilized as a substrate, acetogens must compete with Fe(III) reducers, sulfate reducers, and methanogens. In most anoxic habitats with low amounts of Fe(III) and sulfate, CO2-dependent methanogenesis dominates as the terminal reductive process during the oxidation of organic matter. H2-CO2-dependent methanogenesis (DG0 o = 130 kJ per mole reaction) is more exergonic than H2-CO2-dependent acetogenesis (DG0 o = 95 kJ per mole reaction; Diekert and Wohlfarth 1994b; Fuchs 1994). At the in situ concentrations of reactants and products found in many anoxic environments, methanogenesis may be energetically more favorable than acetogenesis (Dolfing 1988). H2-thresholds (i.e., the minimum concentration of H2 required for the uptake of H2 by a cell) decrease with increasing redox potential of the energy-yielding reaction (Conrad 1996). The redox potential of the CO2/acetate half-cell reaction is 290 mV and is more negative than that of most other terminal-electronaccepting processes. Thus, under pure culture conditions, the H2 threshold of acetogens is higher than that of other hydrogenotrophic anaerobic bacteria (Cord-Ruwisch et al. 1988). For example, the H2 threshold for acetogens when CO2 is utilized as a terminal electron acceptor (362–4,660 ppm) is 10- to 100-fold higher than that of methanogens. Thus, on these theoretical grounds, H2-dependent acetogenesis is not a competitive process at low concentrations of H2. The capacity of acetogens to utilize various substrates simultaneously might contribute to their competitiveness in nature. Aromatic methoxyl groups are more readily utilized by deep subsurface acetogens in the presence of H2 (Liu and Suflita 1993). During mixotrophic growth on H2 and lactate, A. woodii can utilize lower concentrations of H2; however, the residual partial pressure is still too high for a successful competition with H2-utilizing methanogens (Peters et al. 1998).

In certain complex microbial habitats (e.g., soils and sediments), the metabolic interactions of anaerobic populations are influenced by unstable physical and chemical parameters such as pH, temperature, periodic oxygenation, spatial arrangements, and different sizes of microbial populations. In certain cases, these complex factors might be favorable for the theoretically disadvantaged acetogens. For example, in freshwater sediments with low pH and low temperature, acetogens can outcompete methanogens for H2 (Phelps and Zeikus 1984; Conrad et al. 1989; Nozhevnikova et al. 1994; Zavarzin et al. 1994; see the section on > ‘‘Diverse Habitats’’ in this chapter). In contrast to laboratory cultures, most in situ microbial habitats are not composed of homogeneous or well-mixed microbial populations. Thus, the success of acetogens to compete for H2 in situ could be enhanced by being spatially associated with H2-producing cells in a heterogenic system. However, information about the in situ spatial distribution of acetogens at the microscale level is scant, mainly because broad-based, groupspecific 16S-rRNA-based probes cannot be developed for the phylogenetically diverse acetogens (see the section on > ‘‘Detection of Acetogens’’ in this chapter). Furthermore, the functional gene probe that has been developed for acetogens appears to be restricted to a subgroup of acetogens and, likewise, is not absolutely specific for acetogens (Lovell and Hui 1991). Methanogens appear to be more sensitive to O2 than acetogens, and acetogens can occur in higher cultured numbers than methanogens in habitats that are subject to fluctuating concentrations of O2 (e.g., soils). In the hindgut of termites, a microbial habitat that can have volumes of 1 ml or less, only the inner portion of the microbe-packed paunch is completely anoxic; the epithelial surface of the paunch is characterized by a diminishing O2 gradient (Brune et al. 1995, 2000). H2-utilizing acetogens from the rumen are more competitive for H2 than H2-utilizing methanogens in certain termite guts (Brauman et al. 1992). The extent to which H2-derived reductant flows toward the reduction of CO2 to acetate rather than the reduction

Acetogenic Prokaryotes

of CO2 to methane varies with the feeding guild of the termite (Brauman et al. 1992). Although acetogens are usually obligate anaerobes, acetogens are relatively tolerant to fluxes of O2 in terrestrial soils (Wagner et al. 1996). The enumerability of acetogenic bacteria in well-drained, oxic soils and freshly fallen litter (Peters and Conrad 1995; Ku¨sel et al. 1999c; Reith et al. 2002), as well as the isolation of S. silvacetica and M. thermoacetica from different soils (Go¨ßner and Drake 1997; Kuhner et al. 1997; Go¨ßner et al. 1999; Karita et al. 2003), attests to the ability of acetogens to cope with fluxes of aeration and to withstand drying under oxic conditions (Wagner et al. 1996). In contrast, the cultured numbers of other so-called obligate anaerobes, like sulfate reducers or methanogens, are negligible in welldrained soils and litter (Peters and Conrad 1995; Ku¨sel et al. 1999c). The ability of acetogens to survive in habitats that are subject to fluctuations in O2 is due in part to their ability to reductively remove traces of O2 (see the section on > ‘‘Tolerance to Oxic Conditions and Metabolism of O2’’ in this chapter). Their survival in such habitats can also be enhanced by trophic interactions with facultative anaerobes. The classic acetogen M. thermoacetica and the fermentative microaerophile Thermicanus aegyptius were co-isolated as a commensal pair from an oxic hightemperature soil (Go¨ßner et al. 1999). The two organisms grow commensally on oligosaccharides (> Fig. 1.18). In addition to the production of fermentation products by T. aegyptius that

Short-chain sugars (e.g., stachyose)

CO2

O2

Succinate Ethanol Acetate

C O M M E N S A L

Thermicanus aegyptius (microaerophile)

changing conditions H2, Formate, Lactate

trophic link

anoxia

Moorella thermoacetica (acetogen)

Acetate

. Fig. 1.18 Scheme illustrating the hypothetical trophic interaction of Thermicanus aegyptius ET-5b and Moorella thermoacetica ET-5a (Modified from Go¨ßner et al. 1999)

I N T E R A C T I O N

1

can be used by the juxtaposed acetogen, the fermentative microaerophile also minimizes the level of incoming O2 in microzones inhabited by acetogens. Collectively, these findings demonstrate that the ecological roles of acetogens are not restricted to anoxic and water-saturated habitats.

Diverse Habitats Acetogens have been mostly obtained from habitats that are permanently anoxic, such as freshwater or marine sediments, sewage sludge, and gastrointestinal tracts (see the section on > ‘‘Description of Species’’ in this chapter). However, during more recent years, the presence and ecological role of acetogens have also been investigated in other habitats, such as hypersaline sediments, deep aquifers, oxic soils, and plant roots. Although the magnitude and consequence of acetogenesis in these diverse habitats are often unclear, the occurrence of acetogens in such diverse habitats underscores the potential importance of acetogenesis at a more global level. Human Colon. The gastrointestinal tracts of mammals are colonized by acetogens (Prins and Lankhorst 1977; Breznak and Kane 1990; Mackie and Bryant 1994; Wolin and Miller 1994; Leedle et al. 1995). In humans, dietary components (cellulose, hemicellulose, pectin, and starch) not absorbed in the upper digestive tract reach the colon where they are fermented by the cooperative metabolism of a great variety of bacterial species (Wolin and Miller 1983). Short-chain fatty acids (acetate, propionate, and butyrate) and gases (CO2 and H2) are the main fermentation products in the colon, and it is estimated that 95 % of the short-chain fatty acids are absorbed and utilized by the host. Part of the daily production of acetate in the intestine (10–30 g of acetate; Royall et al. 1990) can be attributed to the activity of acetogens, as evidenced by the formation of [1,2-14C] acetate from [3,4-14C] glucose in fecal incubations (Miller and Wolin 1996). Part of the H2 formed during colonic fermentation is exhaled in breath and vented in flatus. A large amount of the H2 that is produced in the colon is consumed via the interspecies transfer of H2. Humans (30–50 % of the European population) who harbor large populations of methanogens (108–1010 cultured methanogens per gram dry wt. of feces) exhale CH4 in detectable concentrations (i.e., >1 ppm of CH4; Miller and Wolin 1982; Pochart et al. 1992). In the colon of these humans, methanogenesis is probably the main hydrogenotrophic process. H2-utilizing Methanobrevibacter smithii is believed to be primarily responsible for producing almost all colonic CH4 (Wolin and Miller 1983). Depending on the human sampled, the cultured number of methanogens in human feces range from undetectable to 1010 per g dry wt. of feces. In contrast, the cultured numbers of H2-utilizing sulfate reducers in the feces of both CH4-excreting and non-CH4-excreting humans are similar (107 per g dry wt. of feces; Dore´ et al. 1995). However, cultured numbers of H2-utilizing fecal acetogens of non-CH4excreting humans are higher than those of CH4-excreting humans (107 versus 105H2-utilizing acetogens per gram dry

41

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Acetogenic Prokaryotes

wt. of feces, respectively). A negative correlation between the numbers of acetogens and methanogens also exists in the rumens of lambs and in the feces of rats (Prins and Lankhorst 1977). These correlations in the relative distributions of H2-dependent anaerobes suggest that H2-dependent acetogenesis is important in gastrointestinal ecosystems which have low numbers of methanogens (Dore´ et al. 1995; Bernalier et al. 1996a). The synthesis of [13C]acetate from 13CO2 by human fecal suspensions is supportive evidence of the hypothesis that reductive acetogenesis is a major colonic process of non-CH4-excreting humans (Lajoie et al. 1988). Numerous H2-utilizing acetogens have been isolated from human feces (Wolin and Miller 1993; Bernalier et al. 1996b, c; Kamlage et al. 1997; Leclerc et al. 1997a), and some of these isolates (i.e., R. hydrogenotrophicus and C. coccoides 1410) form a separate subgroup within the clostridial cluster XIVa (Collins et al. 1994). Rumen Ecosystems. The rumen is arguably the most completely described, intensively investigated gastrointestinal ecosystem, and the classic studies of Hungate (1966) and coworkers were formative to our general understanding of gut ecosystems and the various symbiotic relationships of gut microbiota. CH4 is the main reduced end product that is produced in the rumen (Mackie and Bryant 1994). The major metabolic groups of microorganisms involved in the overall decomposition of organic matter in the rumen are described in the frequently cited three-stage model of McInerney and Bryant (1981). The molar proportions of short-chain fatty acids produced in the rumen approximate 63 % acetic, 21 % propionic, 14 % butyric, and 2 % higher acids. Most (60–80 %) of the daily energy needs of the ruminant are provided by absorption of short-chain fatty acids from the rumen. The loss of energy in the end product CH4 approximates 5–15 % of the feed energy consumed by the host animal (Mackie and Bryant 1994). It would theoretically be beneficial to the host animal if the H2 and CO2 that otherwise are consumed in the production of CH4 were instead converted to acetate by acetogens (Wood and Ljungdahl 1991). CH4 is a greenhouse gas, and since ruminants produce approximately 50 % of the global biogenic emission of CH4 (Mackie and Bryant 1994), a reduction in the emission of methane by ruminants would likewise have theoretically positive effects on the global greenhouse gas budget. In general, methanogens are the dominant H2-utilizers in the rumen (Hungate 1976; Bryant 1979). Thus, the addition of a specific inhibitor for methanogens (e.g., bromoethanesulfonic acid) is usually necessary to enrich or isolate H2-utilizing acetogens. The addition of rumen fluid may be stimulatory to the growth of acetogens found in the rumen (Rieu-Lesme et al. 1995). The capacity of acetogens to utilize methanol, sugars, or methoxylated aromatic compounds has been used to isolate or enumerate ruminal acetogens. Depending on the host animal and substrate used, the number of cultured acetogenic bacteria in the rumen range from 106 to 109 acetogens per gram of ruminal content (Sharak Genthner et al. 1981; Krumholz and Bryant 1986; Leedle and Greening 1988). The number of cultured methanogens in the rumen approximates 108–109

methanogens per gram of ruminal content (Leedle and Greening 1988). Postprandial changes in the population profiles of ruminal acetogens and methanogens occur in steers fed either highor low-forage diets. After a shift from a low-forage diet to a higher input of readily available carbohydrate, a twofold increase in the cultured numbers of both H2-oxidizing methanogens and acetogens occurs after 1–2 h of feeding. The cultured numbers of ruminal acetogens obtained from steers maintained on a high-forage diet are higher than the cultured numbers of ruminal methanogens, suggesting that acetogens capable of utilizing H2 grow preferentially on organic substrates in the rumen (Leedle and Greening 1988). Numerous acetogens have been isolated from the rumen (Sharak Genthner et al. 1981; Krumholz and Bryant 1986; Greening and Leedle 1989; Rieu-Lesme et al. 1995; Rieu-Lesme et al. 1996a, 1996b, 1998; see the section on > ‘‘Description of Species’’). However, the diversity and ecology of acetogens and the competition between acetogens and methanogens in ruminal ecosystems remain poorly resolved, and a substantial amount of information in these areas will be needed before the methanogenic nature of the rumen can be reasonably and successfully manipulated. In this regard, inhibitors (i.e., N-substituted derivatives of p-aminobenzoic acid) of the enzyme responsible for the synthesis of methanopterin (an intermediate in methanogenesis) can block the growth of certain methanogens but not interfere with the growth of acetogens (Dumitru et al. 2003). Termite Guts. About 4 % of the plant material synthesized annually in terrestrial ecosystems is consumed by termites (Ljungdahl and Eriksson 1985). Termites can be divided into four different feeding guilds: the wood-, grass-, and soil-feeding, and the fungus-growing termites (Brauman et al. 1992). The digestion of lignocellulose by wood-feeding termites has been studied for over seven decades (e.g., Cleveland 1925; Hungate 1943; Varma et al. 1994). The wood-feeding termites include the so-called ‘‘lower’’ termites, like the well-studied Reticulitermes flavipes, and also some ‘‘higher’’ termites like Nasutitermes nigriceps. All termites harbor a diverse and dense hindgut microbial community that aids in digestion and is the source of fermentation products such as acetate, H2, and CH4. The hindgut microbiota of the wood-feeding lower termites is composed of cellulolytic protozoa and bacteria that symbiotically affect an essentially acetogenic decomposition of wood polysaccharides (Brauman et al. 1992). Acetate constitutes 94–99 % of the short-chain fatty acid pool in the extracellular hindgut fluid of R. flavipes. Protozoa initially convert cellulose to acetate, H2, and CO2 in a 1:2:1 ratio, and acetogenic bacteria subsequently convert the H2 and CO2 to acetate (Breznak and Switzer 1986). Acetate (1) is absorbed in the hindgut, (2) is oxidized by the termite, and (3) can support up to 100 % of the insect’s energy requirement (Breznak 1994). Thus, H2utilizing acetogens outcompete H2-utilizing methanogens in wood-feeding termites; similar patterns occur for grass-feeding termites (Breznak 1994). In contrast, H2-dependent acetogenesis is of little significance in fungus-growing and soilfeeding termites, both of which evolve more methane than do

Acetogenic Prokaryotes

wood- and grass-feeding termites (Brauman et al. 1992). It is not known whether the nature of the food consumed or other parameters, like a modified gut anatomy or digestive physiology, affect the terminal electron flow in the hindgut microbiota. The core of dissected hindguts in R. flavipes is anoxic, whereas the peripheral lumen of dissected hindguts exhibits high oxygen uptake rates, suggesting that the hindgut has an oxic periphery and an anoxic center under in situ conditions (Brune et al. 1995; Ebert and Brune 1997). The influx of oxygen via the gut epithelium and its reduction in the hindgut periphery appears to have a significant impact on the flow of carbon and reductant within the hindgut microbial community (Tholen and Brune 2000). Although acetogenic bacteria are obligate anaerobes, it is now well known that some of them are not only quite tolerant to oxygen exposure but can also reduce oxygen (Ku¨sel et al. 2001; Karnholz et al. 2002; Drake et al. 2002; Boga and Brune 2003; Drake and Ku¨sel 2003). The highest concentration of H2 in the hindgut occurs in the central region in which H2-producing protozoa also occur. The central region of the hindgut is also the major zone of H2 consumption. Excess H2 diffuses radially outward to the gut epithelium where it seems to be consumed by methanogens, which, for unknown reasons, appear to preferentially colonize the region near the gut wall (Leadbetter and Breznak 1996; Ebert and Brune 1997). Spirochetes occur in the central region of the hindgut and are among and often attached to the H2-producing protozoa. Pure cultures of termite gut spirochetes (e.g., ‘‘Treponema primitia’’) catalyze the synthesis of acetate from H2 and CO2 (Leadbetter et al. 1999; Graber and Breznak 2004a; Graber et al. 2004b). Attachment of acetogenic spirochetes to termite gut protozoa yields a syntrophy that is based on the interspecies transfer of H2 to the acetogen; this symbiosis provides H2 concentrations well above the known H2-threshold values for acetogens. In situ activity measurements of acetogenic bacteria in combination with axial H2 profiles in the highly compartmentalized hindgut of soil-feeding higher termites (Cubitermes spp.) revealed that acetogenesis might have a larger impact on the overall carbon flow than expected from previous observations. Acetogens in the posterior hindgut seem to be supported by either substrates other than H2 or by a cross-epithelial H2 transfer from anterior gut regions, which may create microniches favorable for H2-dependent acetogenesis (Schmitt-Wagner and Brune 1999; Tholen and Brune 1999). Thus, the in situ spatial distribution of acetogens and their orientation in metabolic gradients contributes to their ability to successfully compete for H2 in the termite gut. Marine, Estuarine, and Freshwater Sediments. Acetogens are ubiquitous and plentiful in the sediments of aquatic habitats. The first species of the genus Acetobacterium was isolated from an estuarine sediment (Balch et al. 1977), but other Acetobacterium species have been isolated from freshwater and marine sediments, and from waterlogged terrestrial soils, such as subsurface sandstone, tundra wetland soils, and fens (see the section on > ‘‘Description of Species’’ in this chapter). In such habitats, acetogens might have to compete for H2. Theoretically, sulfate reducers and methanogens can maintain H2

1

concentrations at levels lower that that needed for the acetogenic reduction of CO2 (Cord-Ruwisch et al. 1988). However, in certain sediments, mildly acidic conditions or low temperatures appear to favor acetogenesis. Acetogens can outcompete methanogens for H2 at the in situ pH of 6.2 and also at more acidic pH values (Phelps and Zeikus 1984). Clostridium scatologenes SL1 is an H2-utilizing acetogen isolated from acidic freshwater sediments and is capable of growing at pH 4 (Ku¨sel et al. 2000). Sodium-proton antiporters (see the section on > ‘‘Conservation of Energy and Bioenergetics’’ in this chapter) might enable certain acetogens to cope with broad variations in pH (Schink 1994). Acetogens successfully compete with methanogens for H2 at an in situ temperature of 4  C in sediments of Lake Constanz (Conrad et al. 1989). Although the partial pressure of H2 measured in the pore water is too low for the utilization of H2 by pure cultures of acetogens, the actual in situ partial pressure of H2 might be higher for acetogens living in close proximity to H2-producing organisms. Acetogenic bacteria are important during the degradation of organic matter in permanently cold sediments, and acetate serves as the primary substrate for methanogenesis. Hydrogenotrophic methanogens from sediments of Lake Constanz can be enumerated and activated only at incubation temperatures of 20  C or higher (Schulz and Conrad 1996). In tundra wetland soils or sediments polluted with paper-mill wastewater, acetogens can successfully compete with methanogens for H2 and methanol at temperatures below 15  C (Nozhevnikova et al. 1994; Zavarzin et al. 1994). Acetate accumulates during the first phase of activity and is followed by a slow acetoclastic methanogenesis. The isolation of psychrophilic or psychrotrophic acetogens from these habitats and from cold lake sediments (Conrad et al. 1989; Kotsyurbenko et al. 1995, 1996; Simankova et al. 2000; Nozhevnikova et al. 2001) underscores the potential importance of acetogenesis in low temperature ecosystems. Very little information is available on the occurrence and activity of acetogens in marine habitats. Cultured numbers of H2-utilizing acetogens and acetate-utilizing sulfate reducers from a marine sediment approximated 105 and 107 cells per g wet wt. of sediment, respectively (Ku¨sel et al. 1999b). The concentration of sulfate in the pore water of marine sediments can vary owing to seasonal thermal stratification. Owing to the depletion of sulfate in the sediment, the flow of carbon and reductant shifts from sulfate reduction to methanogenesis (Hoehler et al. 1999). During this transition period, the concentration of acetate increases, which might be due to a temporary decoupling of acetate-producing and acetate-consuming processes (Sansone and Martens 1982). The concentration of H2 in the sediment is elevated at the beginning of this transition period, thus making the acetogenic reduction of CO2 more favorable. The production of [14C]-acetate from 14CO2 occurs at rates comparable to those of methanogenesis or sulfate reduction during their respective period of dominance (Hoehler et al. 1999). Thus, the transient acetogenic reduction of CO2 may be important in marine sediments and also in other ecosystems that experience geochemical fluctuations.

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Acetogenic Prokaryotes

Hypersaline Environments. Little is known about the presence of halotolerant or halophilic anaerobes or the anaerobic breakdown of organic matter in hypersaline ecosystems (e.g., inland lake and marine salterns). In hypersaline ecosystems, the salinity ranges from 9 % to over 20 % (wt./vol.) and the pH ranges from 7 to 10 (Ollivier et al. 1994). The concentration of sulfate, which can be an important electron acceptor, varies from 0.5 (Dead Sea, Israel) to 21 g per liter (Soap Lake, Washington State, United States). High salt concentrations prevent the growth of vertebrates, and only invertebrates, algae, and prokaryotes are present when the salinity exceeds 10 % (wt./vol.; Ollivier et al. 1994). The concentrations of volatile fatty acids and H2 present in sediments increase when the concentration of salt is high (Oren 1988), indicating that the anaerobic mineralization of organic matter is inhibited by salinity. Bacteria maintain high intracellular salt concentrations or accumulate organic osmolytes (e.g., betaine) to maintain cell turgor at high salt concentrations (Oren 1999). Betaine does not appear to be a substrate for methanogens or sulfate reducers. However, halophilic acetogens (e.g., A. arabaticum) decompose betaine to trimethylamine and acetate (Zhilina and Zavarzin 1990). Trimethylamine is subsequently utilized by methylotrophic methanogens and, thus, is a trophic link between acetogens and methanogens. Since halophilic acetoclastic methanogens are unknown, and since acetateconsuming sulfate reducers are inhibited at high salinity (Widdel 1988), the anoxic fate of acetate in these habitats is unresolved (Zavarzin et al. 1994). The haloalkaliphilic acetogens N. acetigena (Zhilina et al. 1996) and N. histidinovorans (Zhilina et al. 1998) were isolated from soda-depositing lakes and utilize glutamate, which can also be accumulated as an osmolyte by moderate halophiles. The anaerobic decomposition of osmoregulatory compounds in halophilic communities might therefore be coupled to the activity of Acetohalobium-type and Natroniella-type acetogens. Species of the acetogenic genera Eubacterium, Sporomusa, and Acetobacterium demethylate the osmolytes dimethylsulfoniopropionate and glycine-betaine to methylthiopropionate and dimethylglycine, respectively; however, only the demethylation of glycine-betaine supported growth of the organism (Jansen and Hansen 2001), indicating that certain in situ transformations of osmolytes by acetogens are not coupled to growth. Waterlogged Soils. The flow of carbon and reductant in flooded rice paddy soils occurs mainly under anoxic conditions, and CH4 is the major reduced end product of organic matter breakdown (Conrad 1993). Root exudates and rice straw are important sources of energy and carbon for microbial activity (Chidthaisong et al. 1999). Approximately 80 % of the CH4 formed is derived from acetoclastic methanogenesis, which is proportionally more than the amount of CH4 formed in a normal methanogenic food web (Chin and Conrad 1995). Thus, acetogens may be important to the formation of acetate in such waterlogged habitats. Hydrogenotrophic methanogenesis in flooded paddy soils and tundra wetland soils is inhibited by low temperatures (Conrad et al. 1989; Nozhevnikova et al. 1994; Chin and Conrad 1995), and low

incubation temperatures have been used to select against hydrogenotrophic methanogens during the enrichment of H2-utilizing acetogens from paddy soils (Conrad et al. 1989). Cultured numbers of acetogens in the soil of rice plant microcosms range from 103 to 105 cells per g dry wt. of soil, and most dominant cultured acetogens belong to the genus Sporomusa (Rosencrantz et al. 1999). However, the occurrence of acetogens in the rhizosphere of rice plants is not resolved. Fluorescent in situ hybridization analysis has revealed the occurrence of Archaea on the surfaces of rice roots (Großkopf et al. 1998). Plant Roots. The rhizosphere is an important microhabitat where complex biogeochemical processes occur at accelerated rates. Plant roots exude easily degradable organic compounds that can chemotactically attract microorganisms (Waisel and Agami 1996). Most rhizosphere bacteria are thought to live near the root tips or in the rhizoplane, which is defined as the root surface and outermost cells of the root. However, little is known about the colonization of the rhizosphere and plant roots by anaerobic bacteria (Großkopf et al. 1998; Hines et al. 1999). Although the rhizospheres of salt marsh vegetation and rice plants might be thought of as anoxic, a gradient of O2 is generated around roots via the transport of O2 that is produced by leaf photosynthesis during the day (Gilbert and Frenzel 1995; Revsbech et al. 1999). Thus, obligate anaerobes colonizing such rhizospheres experience periods of elevated O2 tension. Sea grass rhizosphere has higher numbers of acetogenic bacteria than unvegetated soil, and acetogenic O-demethylation activity is tightly associated with sea grass roots (Ku¨sel et al. 1999b). Hybridization of root thin sections with 33P-labeled probes specific for Acetobacterium revealed the intercellular colonization of sea grass roots by Acetobacterium-like bacteria (Ku¨sel et al. 1999b). The Acetobacterium-like bacteria occur mostly in the rhizoplane and outermost cell layers of the cortex. An H2-utilizing acetogen, RD1, was obtained from the highest, growth-positive dilution of a sea grass root most-probablenumber series, and analysis of the 16S rRNA gene sequence indicated that the acetogen was closely related to a bacterium (Clostridium glycolicum; 99.7 % gene sequence similarity) not previously known to be an acetogen (Ku¨sel et al. 2001; > Fig. 1.19). Retrieval of formyltetrahydrofolate synthetase sequences from salt marsh plant roots indicates that such roots are colonized by diverse acetogens most closely related to the genera Sporomusa, Acetobacterium, Clostridium, and Eubacterium (Leaphart et al. 2003). Acetogens associated with salt marsh plant roots might also display a high tolerance to O2 (Ku¨sel et al. 2003). These results indicate that the biogeochemistry of the sea grass rhizosphere fosters the growth of acetogens in a habitat classically considered to be sulfate reducing. Deep Subsurface. Many subterranean environments are anoxic and habitats for anaerobic microorganisms. In the past, deep subsurface microbial communities have been thought to be supported by organic matter deposited with the formation of sediments or by organic matter that migrated from the surface along different flowpaths (Krumholz 2000). Thus, most studies

Acetogenic Prokaryotes

a

1

b

2

. Fig. 1.19 Electron micrographs of (a) the acetogen Clostridium glycolicum RD-1 that was isolated from a sea grass root (used with permission from Ku¨sel et al. 2001) and (b) the nitrogen-fixing soil bacterium Clostridium akagii (Kuhner et al. 2000). These organisms have connecting filaments that might provide cells with a means of remaining in close proximity to one another for either structural or communication purposes under certain in situ conditions. The structural nature of the filaments is not fully resolved, but recent ultra structural analyses suggest that the outer portion of the filament is an extension of the outer surface layer of the cell. Bars are in micrometers

have focused on the occurrence and activity of microorganisms within recently deposited or highly permeable sediments rather than from consolidated subterranean rock. However, diverse microbial communities also occur in subsurface fractured granitic rock, in suboceanic basalts, and in deep sediments of oceans (Krumholz 2000; Kotelnikova 2002). It is estimated that the global carbon content in subsurface prokaryota is comparable to the carbon content stored in terrestrial plants (Whitman et al. 1998). The density of a microbial community is limited by nutrient availability, which decreases in general with increasing depth. In deep, low organic carbon sediments in the Woodland Basin of the Pacific Ocean, numbers of cultured H2-utilizing acetogens decrease with increasing depth from a surface maximum of approximately 106 cells per ml of sediment to negligible numbers at a depth of 800 m below the seafloor (Wellsbury et al. 1997, 2002). However, microbial life might also be supported by other mechanisms, e.g., by the use of rock- or sediment-bound organic material previously thought to be unavailable. Organic matter trapped in shales during deposition in the Cretaceous period (about 100 million years ago) can fuel heterotrophic microbial communities and the formation of acetate in adjacent permeable sandstones (Krumholz et al. 1997). From these sandstones, the acetogen A. psammolithicum was isolated with H2-CO2 as a substrate (Krumholz et al. 1999). However, the reduction of sulfate might be the dominant sink for H2-derived reductant, and organic compounds might be the main substrates for acetogens in such terrestrial subsurface habitats (Krumholz et al. 1999). In suboceanic sediments, bacterial populations and their activities can increase even in deeper layers near gas hydrate

1

zones. Pore water concentrations of acetate can reach surprisingly high concentrations of approximately 15 mM at 700 m below the seafloor, approximately 100 times higher than average near surface concentrations (Wellsbury et al. 1997, 2000). Acetate seems to be the principal energy source for methane formation, and high turnover rates of acetate indicate an upward migration of high concentrations of dissolved organic carbon into the sediments. Apparently, the bioavailability of sedimentary organic matter appears to be enhanced by low-temperature heating during burial (Wellsbury et al. 1997). In addition to this biological enhanced formation of acetate, acetate can also be formed by thermogenic alteration of organic matter at temperatures above 80 C. The potential role of mesophilic and/or themophilic acetogens in the formation of acetate in these deep sediment layers is unclear. Dissolved organic carbon may serve as a source of energy and carbon for microorganisms in deep subterranean groundwater of thin granitic fractures (Kotelnikova 2002). In addition, concentrations of dissolved H2 in these deep aquifers can be significantly higher (20–100 mM) than in other aquatic surface habitats (Krumholz 2000). Acetogenic and acetotrophic methanogenic bacteria dominate the viable cell counts of different physiological groups in deep granitic groundwater that contains H2 and CH4 (Kotelnikova and Petersen 1997; Kotelnikova and Pedersen 1998). The cultured numbers of H2-utilizing acetogens approximate up to 104 cells per ml of groundwater. In microcosms containing granitic groundwater, 14CH4- and 14 C-labeled acetate are formed from 14CO2, and 14C-labeled acetate is converted to 14CH4. Thus, chemolithoautotrophic microorganisms that can grow on H2-CO2 might act as primary producers of organic carbon, initiating heterotrophic food chains in deep subterranean habitats. Acetogenesis appears to be involved in these H2-based autotrophic biospheres (Haveman and Pedersen 2002; Kotelnikova 2002). The origin of H2 in subsurface habitats appears to be diverse. H2 may result from mixed geochemical and biogenic reactions including the hydrolysis of water under strongly reduced conditions by ferrous iron present in basalt (Kotelnikova 2002). In controlled laboratory experiments, basalts as well as granitic rock samples incubated with buffered water produce H2 gas (Stevens and McKinley 1995). However, whether those levels of H2 could be produced at environmentally relevant pH values or sustained over geological time has been questioned (Anderson et al. 1998). Nonetheless, deep basalt aquifers can contain up to 60 mM dissolved H2, and autotrophs outnumber heterotrophs in such habitats. Stable isotope measurements suggest that autotrophic methanogenesis dominates this lithoautotrophic microbial ecosystem in deep basalt aquifers (Stevens and McKinley 1995). Oxic Soils. Anoxic microzones can occur in oxic soils and litter when the consumption of O2 exceeds its supply (Tiedje et al. 1984; Sexstone et al. 1985; Smith and Arah 1986; Van der Lee 1999). Forest, agricultural, and grassland soils have a tremendous capacity to form acetate from endogenous organic matter under anoxic conditions (up to 15 g of C-acetate per kg

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dry wt. of soil; Ku¨sel and Drake 1994; Peters and Conrad 1996; Wagner et al. 1996). Beech leaf litter and spruce litter display a spontaneous capacity to form aliphatic acids (mainly acetate), alcohols, H2, and CO2 under anoxic conditions, indicating that a subcommunity of the microbiota can respond rapidly to anoxic conditions (Ku¨sel and Drake 1996; Reith et al. 2002). Supplemental H2, CO, or ethanol is converted to acetate by soils under anoxic conditions in stoichiometries that approximate those associated with H2-, CO-, or ethanol-dependent acetogenesis. The acetogen S. silvacetica was isolated from forest soil and utilizes H2 and ethanol (Kuhner et al. 1997). Acetogenic activities of soils are relatively stable when soils are subjected to oxic drying or fluxes of O2 (Wagner et al. 1996). At in situ temperatures and independent of moisture content and the concentrations of acetate that are formed, acetate is a stable end product in anoxic soil and litter microcosms. After extended incubation periods (1–3 months), acetoclastic methanogenesis is induced (Ku¨sel and Drake 1995), which is consistent with (1) the view that oxic soils are not a significant source of methane (Boone 1991; Tyler 1991) and (2) the fact that the number of cultured methanogens is negligible in oxic soils (Peters and Conrad 1995; Ku¨sel et al. 1999c). The cultured number of anaerobes from both forest mineral soil and litter is identical with the cultured number of acetateproducing anaerobes (Ku¨sel et al. 1999c). H2-utilizing acetogens are a dominant group of the cultured anaerobes and approximate 104 to 105 cells per g of dry soil or litter (Peters and Conrad 1995; Ku¨sel et al. 1999c). Because H2 is thermodynamically less than an ideal substrate for acetogens, i.e., for reducing CO2 to acetate (see the section on > ‘‘Use of Diverse Electron Acceptors’’ in this chapter), soil acetogens capable of utilizing organic molecules are likely to occur in greater numbers than those that respond to H2 under laboratory conditions; thus, H2 likely reveals only a small subset of the total acetogenic population. In addition to acetogens, other acetate-forming microorganisms (e.g., facultative members of the Enterobacteriacae) are plentiful in soils and are likely responsible for the majority of acetate formed anaerobically in soils (Ku¨sel 1999c). Independent of these considerations, only 1 % of the cultured H2-utilizing soil acetogens are detected after pasteurization (Ku¨sel et al. 1999c), indicating that (1) a large percentage of the sporeforming soil acetogens are in a vegetative active state or (2) the dominant acetogens present in soil and litter are not sporeformers. The capacity of soils to form acetate from H2-CO2 is enhanced by elevated temperatures (e.g., 30  C and 55  C; Ku¨sel and Drake 1995; Wagner et al. 1996), suggesting that high temperature soils harbor thermophilic acetogens. The isolation of different strains of M. thermoacetica from soils from Kansas, Egypt, and Japan demonstrates that this classic acetogenic thermophile has a wide geographical distribution in oxic soils (Go¨ßner and Drake 1997; Go¨ßner et al. 1998, 1999; Karita et al. 2003). Because of the apparent stability of acetate under anoxic conditions, and because of the temporal and spatial variability of O2 in soils, the consumption of acetate that is formed anaerobically might be linked to oxidative processes. O2 is rapidly

consumed when it is added to litter or soil that has been incubated under anoxic conditions (Ku¨sel and Drake 1995; Wagner et al. 1996; Ku¨sel et al. 1999c), and the consumption of O2 is concomitant to an increase in CO2 and the disappearance of anaerobically formed acetate according to the following stoichiometry: CH3 COOH þ 2O2 ! 2CO2 þ 2H2 O

ð1:14Þ

The rate at which acetate is consumed exceeds the rate at which acetate is formed, indicating that acetate undergoes a rapid in situ turnover. The turnover of acetate can also be linked to other oxidative processes in soils, such as denitrification (Ku¨sel and Drake 1995; Wagner et al. 1996) or Fe(III) reduction (Ku¨sel et al. 2002). Thus, under in situ conditions, the acetate formed in anoxic microzones of oxic soils is likely subject to rapid consumption via (1) the diffusion of O2 into formerly anoxic zones or (2) the transport of acetate with the soil solution into zones where electron acceptors like O2, nitrate, or Fe(III) are present (> Fig. 1.20). These findings indicate that acetate is a trophic link between the different anaerobic and aerobic microbial populations that collectively decompose organic matter in oxic soils.

Organic Carbon in Soil Solution oxic surface hydrolysis, decomposition, & oxidative processes

CO2

[Intermediates] acetate formation via fermentation & acetogenesis

methanogenesis during periods of water saturation

CH4

Acetate NO3− Fe(III)

N2, N2O, NH4+

Fe(II)

O2

anoxic core

H2O increasing anoxic conditions

CO2

. Fig. 1.20 Cross section of a soil aggregate showing a hypothetical anoxic core and possible trophic links between acetate and other redox processes during the oxidation of soil organic carbon to CO2 (Modified from Drake et al. 1997)

Acetogenic Prokaryotes

Biotechnological Applications of Acetogens The biotechnological application of acetogens has been the subject of numerous investigations. However, to date, a commercial-scale application of an acetogen or acetogenesis has not been reported. It is beyond the scope of this chapter to evaluate this topic in detail, and the reader is directed to reviews for further information (Wiegel 1990, 1994; Lowe et al. 1993).

Commercial Production of Acetic Acid from Sugars Acetogens convert sugars stoichiometrically to acetate. This metabolic capacity has been the main focal point of studies designed to evaluate the commercial application of acetogens. Acetic acid is produced commercially from feedstock compounds (e.g., methanol), and global production in 2001 approximated 1010 kg (Anonymous 2002; Causey et al. 2003). Acetic acid can be produced microbiologically from sugars. In the twostage vinegar process, a hexose is converted to two molecules of acetic acid by the sequential activities of a yeast (e.g., Saccharomyces cerevisiae) that anaerobically produces two molecules of ethanol per hexose and of an aerobe (e.g., A. aceti) that only partially oxidizes ethanol to acetic acid. In contrast, the production of acetic acid by acetogenic bacteria is a single-stage process. Although acetogenesis conserves all of the carbon of glucose in the product acetic acid and might therefore be considered the ideal microbial process for the commercial production of acetic acid, the commercialization of the process has thus far not been realized. Furthermore, both microbiological processes are about 35 % more expensive than the cost of the synthetic process from feedstock chemicals (estimated at $0.30/lb of acetic acid in 1991; Busche 1991). Escherichia coli has been genetically modified to produce 2 moles of acetate per mole glucose fermented (Causey et al. 2003); such an organism might be competitive with the two-stage vinegar process. The two main unsolved problems for the commercialization of the acetogenic process are as follows: (1) Acetogens are inhibited by high concentrations of acetate and (2) acetogens do not grow under acidic conditions. No known acetogen can adequately produce acetic acid at the concentrations required (i.e., 50 g of acetic acid per liter) for the process to be commercially feasible (Wiegel 1994). These problems have been addressed in numerous studies, but significant breakthroughs in overcoming these problems have not been reported (Schwartz and Keller 1982; Wang and Wang 1983, 1984; Ljungdahl et al. 1985, 1989; Sugaya et al. 1986; Klemps et al. 1987; Brumm 1988; Von Eysmondt et al. 1990; Ibba and Fynn 1991; Parekh and Cheryan 1991; Cheryan and Parekh 1992). Commercialization of acetogenesis would theoretically become cost-competitive with the synthetic process if an acetogenic bacterium that could tolerate acidic conditions and produce high concentrations of acetic acid were either discovered or engineered (Busche 1991). Although mutants have been obtained that have increased tolerance to acidic acid and acidic conditions, such mutants grow poorly

1

(Schwartz and Keller 1982; Wiegel 1994). The strong uncoupling effect of acetic acid and protons on the proton motive force (DpH) and transmembrane electrical potentials (DC) of acetogens results in a collapse of the cell’s ability to conserve energy and, thus, makes it unlikely that simple mutations could circumvent these problems and yield a mutant with tolerance to acetic acid and acidic conditions that is grossly different from the parent strain’s tolerance (Baronofsky et al. 1984). Although sugar dimers can be utilized by certain acetogens (e.g., cellobiose is a substrate for R. productus; Lorowitz and Bryant 1984), a somewhat serious disadvantage of commercializing acetogenesis is that acetogens do not degrade large sugar polymers (e.g., cellulose). However, two acetogens, ‘‘B. formatexigens’’ (Wolin et al. 2003) and M. thermoacetica strain F21 (Karita et al. 2003), with the capacity to degrade cellulose have recently been found. In addition, cocultures of cellulolytic Clostridium thermocellum and the thermophilic acetogen T. kivui convert cellulose to acetate, with near full recovery of cellulose-derived carbon in acetate (Le Ruyet et al. 1984). Similar results were obtained with a coculture of a cellulolytic strain of Ruminococcus albus and the unclassified acetogen HA (Miller and Wolin 1995). Ruminococcus albus forms ethanol and H2 as reduced end products in the absence of the acetogen HA, but in coculture with HA, the reducing equivalents derived from the oxidation of cellulose-derived hexoses are utilized by the acetogen via the interspecies transfer of H2. These findings suggest that acetogens can in fact be utilized for the conversion of cellulolytic material to acetate. Clostridium lentocellum strain SG6 forms high amounts of acetate from cellulose (Ravinder et al. 2001). Although the metabolism of this strain was described as acetogenic, significant amounts of ethanol are also produced and the engagement of the acetylCoA pathway during the degradation of cellulose is therefore uncertain. Despite the limitations and cost barriers outlined above, the potential use of calcium-magnesium acetate as an environmentally safe road deicer and in controlling sulfur emissions during the combustion of high-sulfur coal has continued to foster interest in the commercial production of acetic acid by acetogens (Ljungdahl et al. 1985, 1989; Wiegel et al. 1990; Parekh and Cheryan 1991; Cheryan and Parekh 1992; Wiegel 1994; Cheryan et al. 1997). Thermophilic species of acetogens (e.g., M. thermoacetica and T. kivuii) offer several theoretical advantages for the commercial production of environmentally safe calcium-magnesium acetate: (1) thermophilic acetogenesis bypasses the need to use sterilized medium, the production of which is costly, (2) the dispersal of thermophiles or their spores in low temperature climates does not constitute an environmental threat, and (3) the growth of pathogens and their subsequent dispersal with the calcium-magnesium acetate would be highly unlikely owing to the thermophilic production conditions (Wiegel 1994). Although T. kivuii displays one of the shortest doubling times of all known acetogens [approx. 2.5 h on either H2-CO2 or glucose (Leigh et al. 1981; Daniel et al. 1990)], it is more sensitive to acetic acid than certain strains of M. thermoacetica or M. thermoautotrophica and, therefore,

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does not appear to be a model of choice for commercial purposes (Wiegel 1994). Moorella thermoacetica and Clostridium thermolacticum can convert lactose to acetic acid when the two organisms are immobilized together in a fibrous-bed reactor (Talabardon et al. 2000). Under these conditions, C. thermolacticum forms lactate, which is subsequently used for lactate-dependent acetogenesis by M. thermoacetica. This trophic association mimics that exemplified by M. thermoacetica and T. aegyptius (Go¨ßner et al. 1999; > Fig. 1.20). Immobilization of acetogens with metabolic partners might offer certain advantages for commercializing the use of acetogens. One approach to increasing the amount of acetate produced by acetogens is to uncouple growth from acetogenesis. Harmaline, a putative inhibitor of Na+/H+ antiporters, uncouples acetogenesis from the growth of A. kivuii (Yang and Drake 1990). In the presence of harmaline, the acetate-to-biomass ratio during H2-dependent acetogenesis increased 13-fold. Thus, use of agents that uncouple growth from the production of acetate might be of value in making acetogenesis more commercially feasible.

Bioconversion of Synthesis Gas to Acetic Acid, Ethanol, and Other Chemicals Synthesis gas is obtained by the indirect liquefaction of coal and mainly consists of H2, CO, and CO2. These gaseous molecules can be converted to acetate by acetogenic bacteria, and the potential use of acetogens for the bioconversion of synthesis gas to acetic acid has been evaluated (Grethlein and Jain 1992). Butyrate and n-butanol are additional products that can be produced from synthesis gas by ‘‘B. methylotrophicum’’ (Worden et al. 1989; Grethlein et al. 1991). Likewise, metabolically altered or unique strains of acetogens (e.g., C. ljungdahlii) can produce ethanol from the components of synthesis gas (Buschhorn et al. 1989; Grethlein and Jain 1992; Tanner et al. 1993; Phillips et al. 1994). Electrochemical processes for converting CO2 to acetate with enzymes from M. thermoacetica have been reported (Shin et al. 2001).

Bioremediation, Bioreactors, and Landfills Although acetogens have robust metabolic capabilities and might be thought of as having significant bioremediation potentials, such potentials have not been extensively examined. Certain acetogens (e.g., ‘‘A. dehalogenans’’) can dehalogenate toxic compounds (Egli et al. 1988; Freedman and Gossett 1991; Traunecker et al. 1991; Meßmer et al. 1996). However, few studies have addressed this potential. On the basis of information to date, the ability to degrade aromatic rings is not a widespread metabolic potential of acetogens. A noted exception is H. foetida (Liesack et al. 1994). Acetyl-CoA synthase can transform 2,4,6-trinitrotoluene (TNT), a highly explosive anthropogenic compound that contaminates certain soils (Preuss et al. 1993; Huang et al. 2000); however, a commercial

(or environmental) application of the potential of acetogens to transform TNT has not been reported. Moorella thermoacetica has been shown to be effective in sequestering (i.e., precipitating) the heavy metal cadmium, suggesting that acetogens might be of applied value in the cleanup of environments or materials contaminated with heavy metals (Cunningham and Lundie 1993). By virtue of their ability to oxidize and consume CO, acetogens have been cited as being significant in the detoxification of environmental CO (Ragsdale 1991). However, the detoxification of environmental CO occurs mainly by abiotic processes in the atmosphere or aerobic CO-oxidizers (e.g., soil carboxydotrophs; Meyer 1988; Meyer et al. 1993). The acetyl-CoA synthase of acetogens transforms 2,4,6-trinitrotoluene (TNT; Preuss et al. 1993; Huang et al. 2000); application of this catalytic potential has not been reported. In methanogenic bioreactors and landfills, acetate that is produced by acetogens is a substrate for acetoclastic methanogens. Thus, acetogens contribute significantly to the turnover of organic matter in methanogenic bioreactors and landfills (McInerney and Bryant 1981; Ibba and Fynn 1991; Wiegel 1994; Barlaz 1997).

Other Potential Applications The commercial production of corrinoids (i.e., vitamin B12) and cysteine by acetogens has been evaluated; however, commercialscale production has not been reported (Koesnandar et al. 1991; Inoue et al. 1992; Lebloas et al. 1994). Acetate kinase from M. thermoacetica is very stable, and its industrial use in the immobilized form has been patented in Japan (Wiegel 1994). A variety of fine chemicals (e.g., enantiomers of malic acid) have been produced with C. formicoaceticum (Eck and Simon 1994a; Eck and Simon 1994b). Moorella thermoacetica has been utilized for the electromicrobial regeneration of pyridine nucleotides (Schulz et al. 1995; see also Gu¨nther et al. 2000). Enhancement of acetogenesis in the rumen might enhance the cost-efficiency of ruminant husbandry and also decrease the emission of the greenhouse gas methane by ruminants (this topic is discussed in the section on > ‘‘Diverse Habitats’’).

Summary and Conclusions The following items summarize the main characteristics of acetogens and the acetyl-CoA pathway: 1. Acetogens belong to the domain Bacteria and use the acetyl-CoA ‘‘Wood/Ljungdahl’’ pathway as a terminalelectron-accepting process. 2. The acetyl-CoA pathway fixes CO2, conserves energy, and produces acetyl-CoA that is utilized in the synthesis of either acetate or biomass. 3. Pathways that are biochemically very similar to the acetylCoA pathway are used by methanogens and sulfate reducers

Acetogenic Prokaryotes

4.

5.

6. 7. 8.

9. 10.

11.

for the oxidation of acetate or the autotrophic fixation of CO2 and the synthesis of biomass. The collective use of acetyl-CoA-synthase-dependent pathways by acetogens, methanogens, and sulfate reducers facilitates an enormous turnover of carbon in the global carbon cycle. An acetylCoA-synthase-dependent process may have been the first autotrophic process on Earth. Acetogens can grow autotrophically and heterotrophically and can oxidize a wide range of carbonaceous substrates, including aromatic compounds and low-molecular-weight halogenated compounds. Two recent isolates are cellulolytic. Twenty-one genera of acetogens have been isolated from very diverse habitats, ranging from the gastrointestinal tracts of mammals and insects to sea grass rhizospheres. Their closest 16S rRNA phylogenetic neighbor is very often not an acetogen, making it impossible to develop a broadbased, 16S rRNA acetogen probe. Although acetogens have been classically regarded as obligate anaerobes, they can tolerate and reduce small quantities of O2 and exist in habitats subject to transient fluxes of O2. Acetogens utilize substrate-level phosphorylation, membranous electron transport systems, and ATPases to conserve energy. Although the acetyl-CoA pathway is the hallmark of acetogens, they can also utilize other terminalelectron-accepting, energy-conserving processes, including the dissimilation of aromatic acrylates and nitrate. Thus, acetogens are not strictly dependent upon acetogenesis. The acetyl-CoA pathway can be repressed when alternative terminal electron acceptors are utilized. Thus, acetogens do not always produce acetate. Acetogens form different types of trophic interactions for other microorganisms, including syntrophic relationships with other anaerobes and commensal relationships with fermentative microaerophiles. Numerous theoretical commercial applications for acetogens have been evaluated; however, to date, no commercial-scale application has been reported.

Numerous unresolved questions might be considered worthy of further investigation: What is the biochemical nature of the diverse catabolic processes of acetogens? How are these processes regulated at the gene level? Are acetogens more capable of degrading cellulose and other large biopolymers than previously thought? Can acetogens oxidize inorganic compounds, or is their catabolism restricted to the oxidation of carbonaceous substrates? What is the in situ impact of acetogens in complex habitats, such as the endorhizosphere of plants, gastrointestinal ecosystems, and the terrestrial subsurface? Can the metabolic capabilities of acetogens be successfully harnessed and put to commercial use? The body of published information on acetogens has reached enormous proportions, and it has not been possible to provide adequate coverage to all of these works in this chapter.

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The authors invite individuals working in this area to send their latest results to the corresponding author (H. L. Drake) so that this information can be included when this chapter is updated at a future date.

Dedication and Acknowledgment This tapestry is dedicated to Harland G. Wood and Lars G. Ljungdahl, the two individuals who carried the ball when no one else could. The authors express their appreciation to Anita Go¨ßner for her many years of excellence in culturing and analyzing acetogens, to Marcus Horn for assistance with the phylogenetic analyses, to Georg Acker for electron microscopy of isolates, to Millie Wood for permission to publish the photo of Harland Wood, to Volker Mu¨ller for helpful discussions on bioenergetics, and to John Breznak, Paul Lindahl, Terry Miller, Steve Ragsdale, and Meyer Wolin for providing unpublished information and helpful suggestions. Current support for the authors’ laboratory is derived in part from funds from the German Research Society (DFG) and the German Ministry of Education, Research, and Technology (BMBF), which is gratefully acknowledged.

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Gilbert B, Frenzel P (1995) Methanotrophic bacteria in the rhizosphere of rice microcosms and their effect on porewater methane concentration and methane emission. Biol Fertil Soils 20:93–100 Go¨ßner A, Drake HL (1997) Characterization of a new thermophilic acetogen (PT-1) isolated from aggregated Kansas prairie soil. Abstr Ann Meet Am Soc Microbiol Abstr N-122:401 Go¨ßner A, Daniel SL, Drake HL (1994) Acetogenesis coupled to the oxidation of aromatic aldehyde groups. Arch Microbiol 161:126–131 Go¨ßner AS, Kuesel K, Devereux R, Drake HL (1998) Occurrence of thermophilic acetogens in Egyptian soils. Abstr Ann Meet Am Soc Microbiol Abstr N-1:366 Go¨ßner A, Devereux R, Ohnemu¨ller N, Acker G, Stackebrandt E, Drake HL (1999) Thermicanus aegyptius gen. nov., sp. nov., isolated from oxic soil, a facultative microaerophile that grows commensally with the thermophilic acetogen Moorella thermoacetica. Appl Environ Microbiol 65:5124–5133 Gottschalk G, Braun M (1981) Revival of the name Clostridium aceticum. Int J Syst Bacteriol 31:476 Graber JR, Breznak J (2004) Physiology and nutrition of Treponema primitia, an H2-CO2-acetogenic spirochete from termite hindguts. Appl Environ Microbiol 70:1307–1314 Graber JR, Leadbetter JR, Breznak J (2004) Description of Treponema azotonutricium sp. nov., and Treponema primitia sp. nov., the first spirochetes isolated from termite guts. Appl Environ Microbiol 70:1315–1320 Grahame DA (2003) Acetate C-C bond formation and decomposition in the anaerobic world: the structure of a central enzyme and its key active-site metal cluster. Trends Biochem Sci 28:221–224 Greening RC, Leedle JAZ (1989) Enrichment and isolation of Acetitomaculum ruminis, gen. nov., sp. nov.: acetogenic bacteria from the bovine rumen. Arch Microbiol 151:399–406 Grethlein AJ, Jain MK (1992) Bioprocessing of coal-derived synthesis gases by anaerobic bacteria. TIBTECH 10:418–423 Grethlein AJ, Worden RM, Jain MK, Datta R (1991) Evidence for production of n-butanol from carbon monoxide by Butyribacterium methylotrophicum. J Ferment Bioengin 72:58–60 Großkopf R, Stubner S, Liesack W (1998) Novel euryarchaeotal lineages detected on rice roots and in the anoxic bulk soil of flooded rice microcosms. Appl Environ Microbiol 64:4983–4989 Gunsalus RP, Romesser JA, Wolfe RS (1978) Preparation of coenzyme M analogs and their activity in the methyl-coenzyme M reductase in Methanobacterium thermoautotrophicum. Biochemistry 17:2374–2377 Gu¨nther H, Walter K, Ko¨hler P, Simon H (2000) On a new artificial mediator accepting NADP(H) oxidoreductase from Clostridium thermoaceticum. J Biotechnol 83:253–267 Ha¨ggblom MM, Berman MH, Frazer AC, Young LY (1993) Anaerobic O-demethylation of chlorinated guaiacols by Acetobacterium woodii and Eubacterium limosum. Biodegradation 4:107–114 Hall IC, O’Toole E (1935) Intestinal florain newborn infants with a description of a new patogenic anaerobe, Bacillus difficilis. Am J Dis Child 49:390–402 Hansen B, Bokranz M, Scho¨nheit P, Kro¨ger A (1988) ATP formation coupled to caffeate reduction by H2 in Acetobacterium woodii Nzva16. Arch Microbiol 150:447–451 Harriott OT, Frazer AC (1997) Enumeration of acetogens by a colorimetric most-probable-number assay. Appl Environ Microbiol 63:296–300 Hashsham SA, Freedman DL (1999) Enhanced biotransformation of carbon tetrachloride by Acetobacterium woodii upon addition of hydroxocobalamin and fructose. Appl Environ Microbiol 65:4537–4542 Hattori S, Kamagata Y, Hanada S, Shoun H (2000) Thermoacetogenium phaeum gen. nov., sp. nov., a strictly anaerobic, thermophilic, syntrophic acetateoxidizing bacterium. Int J Syst Evol Microbiol 50:1601–1609 Haveman SA, Pedersen K (2002) Distribution of culturable microorganisms in Fennoscandian Shield groundwater. FEMS Microbiol Ecol 39:129–137 Heijthuijsen JHFG, Hansen TA (1986) Interspecies hydrogen transfer in cocultures of methanol-utilizing acidogens and sulfate-reducing or methanogenic bacteria. FEMS Microbiol Ecol 38:57–64 Heijthuijsen JHFG, Hansen TA (1989) Selection of sulphur sources for the growth of Butyribacterium methylotrophicum and Acetobacterium woodii. Appl Microbiol Biotechnol 32:186–192

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2 Virulence Strategies of Plant Pathogenic Bacteria Maeli Melotto1 . Barbara N. Kunkel2 1 Department of Biology, University of Texas, Arlington, TX, USA 2 Department of Biology, Washington University, St. Louis, MO, USA

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 The Biology of Bacterial Plant Pathogens . . . . . . . . . . . . . . . . . . 62 Virulence Strategies in the Early Stages of Infection . . . . . . . 62 Extracellular Polysaccharides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 65 Plant Cell Wall Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . . 66 Production of Low-Molecular-Weight Phytotoxins . . . . . . . . . 66 Lipodepsipeptide Toxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 Modified Peptide Toxins with Antimetabolite Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 Coronatine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 Syringolin Group . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68 Type III Secretion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Structure and Components of TTSS of Bacterial Plant Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 Identification of Type III Effectors . . . . . . . . . . . . . . . . . . . . . . . 69 Elucidating the Function of Type III Effectors . . . . . . . . . . . 70 Modulation of Plant Hormone Homeostasis and Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 Salicylic Acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 Jasmonates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 Ethylene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 Abscisic Acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 Auxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 The Complexities of Hormone Signaling Networks in Plant-Microbe Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 Challenges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75 Abstract Plant pathogenic bacteria have evolved several unique virulence strategies to successfully infect their hosts. One current area of intense research in the field of plant-pathogen interactions is the identification and characterization of pathogen virulence factors and the elucidation of their mode of action within the host. This chapter summarizes recent progress in this area of research, focusing on four Gram-negative bacterial pathogens that grow

on living tissue and cause primarily leaf spotting or wilt diseases of plants: Pseudomonas syringae, Xanthomonas campestris, Ralstonia solanacearum, and Erwinia amylovora, the causal agents of leaf spots, leaf blights, vascular wilts, and fire blights, respectively. The focus is on these pathogens because significant progress has been made in recent years toward elucidating the molecular mechanisms underlying their virulence. The recently available genome sequence data of various strains of several of these pathogens have also begun to provide additional insight into their virulence strategies. Further, because several of these pathogens can infect Arabidopsis thaliana, use of molecular and genetic approaches to investigate the mode of action of pathogen virulence factors within this host has significantly contributed to our understanding of the virulence strategies of these plant pathogenic bacteria.

Introduction Plant pathogenic bacteria, like bacterial pathogens that infect animals, must be able to evade or suppress general antimicrobial defenses and acquire nutrients and water from their hosts to successfully colonize and grow within host tissue. Plant pathogenic bacteria have adapted well to their hosts, which are structurally and physiologically quite different from animals. Since successful infection relies to a great extent on the ability of a pathogen to modulate the physiology of its host, plant pathogenic bacteria have evolved several unique virulence strategies in addition to virulence mechanisms also utilized by bacterial pathogens of animals. One current area of intense research in the field of plantpathogen interactions is the identification and characterization of pathogen virulence factors and the elucidation of their mode of action within the host. This chapter summarizes recent progress in this area of research, focusing on four gram-negative bacterial pathogens that grow on living tissue and cause primarily leaf spotting or wilt diseases of plants: Pseudomonas syringae, Xanthomonas campestris, Ralstonia solanacearum, and Erwinia amylovora, the causal agents of leaf spots, leaf blights, vascular wilts, and fire blights, respectively (Schroth et al. 1981; Chan and Goodwin 1999; Eastgate 2000; Genin and Boucher 2002). The focus is on these pathogens because significant progress has been made toward elucidating the molecular mechanisms underlying their virulence in recent years (Staskawicz et al. 2001; da Silva et al. 2002; Salanoubat et al. 2002; Buell et al. 2003; Buttner and Bonas 2003; Angot et al. 2006; Bocsanczy et al. 2008; Boch and

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Bonas 2010). The recent genome sequence data made available for various strains of several of these pathogens have also begun to provide additional insight into their virulence strategies (da Silva et al. 2002; Salanoubat et al. 2002; Buell et al. 2003; Feil et al. 2005; Joardar et al. 2005; Smits et al. 2010; for a complete list of bacterial genomes refer to the National Center for Biotechnology Information database at http://www.ncbi.nlm.nih.gov/). Further, as several of these pathogens can infect Arabidopsis thaliana, a widely studied model plant, use of molecular and genetic approaches to investigate the mode of action of pathogen virulence factors within this host has significantly contributed to our understanding of the virulence strategies of these plant pathogenic bacteria (Quirino and Bent 2003; Koornneef and Meinke 2010; Nishimura and Dangl 2010). For reviews on several other fascinating groups of plant-associated microbes, the tumor-inducing Agrobacterium spp., the soft-rot Erwinia species, and the rootnodulating Rhizobia, refer to several recent articles (Gelvin 2003; Toth et al. 2003; Deakin and Broughton 2009; Mallick et al. 2010; Pitzschke and Hirt 2010; Billing 2011).

The Biology of Bacterial Plant Pathogens Phytopathogenic bacteria colonize all plant tissues. Although bacteria can live in the phyllosphere or rhizosphere without causing any harm to the plant, to become a fully virulent pathogen, they must penetrate plant tissues. In many cases, bacterial penetration is a passive process, in which bacteria take advantage of wounds and openings in the plant body or they can be directly deposited by insect vectors. Once inside the plant tissues, different species can inhabit the dead xylem vessels or live in phloem sieve elements; however, the majority of bacterial pathogens are limited to the intercellular space, that is, apoplast (Beattie and Lindow 1994; Alfano and Collmer 1996; Agrios 2005). Although this view may be changing, in general, the apoplast is considered to be a nutrient-poor, unfavorable environment for most microorganisms, containing several antimicrobial compounds (Dangl and Jones 2001; Dixon 2001; Glazebrook 2001; Pignocchi and Foyer 2003; Rico and Preston 2008). Additionally, plant defenses that are induced upon microbial attack are often targeted to the intercellular space (Wang et al. 2005; Bednarek et al. 2010; Wang and Dong 2011). Given the physiology of plants and the nature of the antimicrobial defense responses they deploy, plant bacterial pathogens have evolved a variety of specialized virulence strategies to facilitate colonization of plant tissue. The achievement of this goal relies to a great extent on the ability of plant pathogens to modulate host physiology. As these pathogens are extracellular, they deploy an arsenal of secreted virulence factors to modulate host cell processes from outside plant cells. These include production of protein virulence factors (or effectors) that are delivered directly into the plant cell cytosol via a specialized, type III secretion system (TTSS; Jin et al. 2003a; Block and Alfano 2011; Lindeberg et al. 2012) illustrated in > Fig. 2.1; cell-cell communication through quorum sensing (Quin˜ones et al. 2005; Chatterjee et al. 2007; Barnard and Salmond 2007); production

of low-molecular-weight phytotoxins that are secreted into the apoplast (Bender et al. 1999); exopolysaccharides; and cell wall degrading enzymes. These virulence factors allow the bacteria to evade, overcome, or suppress antimicrobial host defenses and elicit the release of nutrients and water from plant cells to ensure successful colonization of the plant apoplast (> Fig. 2.2). Additionally, some plant pathogens may directly modulate hormone signaling within their hosts through the production of plant hormone analogs (Melotto et al. 2008a; Spoel and Dong 2009; Lee et al. 2009; Grant and Jones 2009; Robert-Seilaniantz et al. 2011; Kazan and Manners 2012). Plant pathogens also express genes believed to help them adapt to the stressful conditions that are constitutively present or that are generated by the host in response to microbial attack. These include the production of proteins and enzymes to counter oxidative stress (e.g., glutathione S-transferase, superoxide dismutase, and catalase), as well as enzymes that may detoxify antimicrobial compounds (Boch et al. 2002; Salanoubat et al. 2002; Buell et al. 2003). Recent studies assessing in planta changes in the transcriptome of X. oryzae pv. oryzicola revealed regulation of genes encoding proteins involved in secretion and transport, tissue adherence, cell wall degradation, and virulence (Soto-Sua´rez et al. 2010). Interestingly, plant-inducible genes in plant pathogens, such as X. oryzae, and plant growth-promoting rhizobacteria, such as P. fluorescens SBW25, fall into similar categories. For instance, P. fluorescens has a functional type III secretion system expressed in the sugar beet rhizosphere (Preston et al. 2001). However, plant-inducible genes in P. fluorescence may be strain-specific indicating that diverse ecological adaptation exists in the pseudomonad group and offering a unique opportunity to address questions about the evolution of plant-microbe interactions (Silby et al. 2009, 2011). In the majority of pathogenic interactions, disease symptoms ensue only after the pathogen has colonized and grown to high levels in the infected tissue. In many cases, disease symptom production is believed to facilitate pathogen release from infected tissue and spread to uninfected tissues and neighboring plants (Agrios 2005). Therefore, the elicitation of disease symptoms is also often considered an important virulence strategy.

Virulence Strategies in the Early Stages of Infection The phyllosphere may be a very harsh environment for pathogens. The leaf surface, in particular, is regularly exposed to extreme conditions such as lack of moisture, ultraviolet irradiation, strong winds, and heat. Nonetheless, bacteria, the most abundant organism on the leaf surface (Lindow and Brandl 2003) can reach high population density (106–107 cells/cm2 of leaf; Andrews and Harris 2000). Pathogens arriving to the leaf surface may have to either quickly penetrate the leaf or express traits that ensure survival until environmental conditions are favorable to penetration. To gain entry into the leaf, bacterial pathogens need to overcome the plants’ active defense against penetration, for example, the closure of the stomatal pore as part

Virulence Strategies of Plant Pathogenic Bacteria

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. Fig. 2.1 Activities of bacterial type three secretion effector proteins injected into the host cell. As extracellular pathogens, bacterial pathogens deploy a variety of secreted virulence factors to modulate the biology of their host cells. These factors include protein virulence factors (or ‘‘effectors’’—labeled with an E in the diagram) that are delivered directly into the plant cell cytosol via the specialized type III secretion system (TTSS). Type III-delivered effectors interact with their targets (labeled with a T) in the host cell and modulate host cell physiology by either inhibiting plant immunity components (currently identified components are depicted in the light green box to the right) or altering hormone homeostasis or signaling. Additionally, some effectors enter the nucleus and function as transcription factors (e.g., TAL-like effectors) upregulating the transcription of susceptibility genes. Ultimately, the collective activity of the effectors promotes pathogen growth and disease symptom development. Disease symptoms facilitate pathogen release from infected tissue, and hence pathogen transmission

of their innate immune system (Melotto et al. 2006; Ali et al. 2007; Zhang et al. 2008; Gudesblat et al. 2009). Certain bacterial pathogens such as X. campestris pv. campestris (Gudesblat et al. 2009), P. syringae pv. syringae (Pss) B728a (Schellenberg et al. 2010), and P. syringae pvs. tabaci, tomato, and maculicola (Melotto et al. 2006) produce phytotoxins that promote pathogenesis by overcoming stomatal immunity as discussed below. Survival as epiphytes may also be considered as a virulence strategy. The mechanism by which bacterial pathogens avoid and/or tolerate stress in the phyllosphere is poorly understood. In an attempt to uncover metabolic activities carried out by bacterial cells when they come in contact with the leaf surface,

Marco et al. (2005) assessed the expression of plant-inducible genes using the pathosystem P. syringae pv. syringae B728a and common bean. This study revealed that genes involved in virulence, transcription regulation, transport, and nutrient acquisition are upregulated in epiphytic bacterial populations. In addition, genes of unknown function were also regulated in epiphytes, suggesting new virulence-associated cellular functions yet to be discovered. Recent studies have revealed that bacterial cell-cell communication through diffusible N-acyl homoserine lactones (AHL), which are quorum sensing signals, occurs in epiphytic bacterial aggregates and controls epiphytic fitness, exopolysaccharide

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. Fig. 2.2 Virulence factors secreted by bacterial plant pathogens. Extracellular bacterial phytopathogens colonize the leaf surface and the intercellular space (i.e., apoplast). These organisms deploy an arsenal of secreted virulence factors to modulate host cell processes from outside plant cells. These factors include (1) a type III secretion system (depicted in > Fig. 2.1), (2) extracellular polysaccharides (EPS) and quorum sensing (QS) signals that may increase pathogen fitness and survival as epiphytes, (3) cell wall degrading enzymes (CWDE) that are secreted through a sec-dependent type II secretion system (Alfano and Collmer 1996; Sandkvist 2001) and function to degrade or remodel the plant cell wall, and (4) low molecular weight toxins (labeled with T) that are secreted into the apoplast, many of which presumably enter or are taken up by plant cells (indicated by the questions mark in the diagram). At least two phytotoxins (coronatine and syringolin A) target stomatal immunity (inhibit stomatal closure) and thus facilitate pathogen entry into the apoplast. In addition, coronatine, a mimic of the plant hormone JA-Ile, is perceived by the COI1/JAZ receptor complex (labeled with R) activating jasmonate signaling and leading to suppression of SA-mediated defenses. Some phytopathogenic bacteria produce syringomycins and syringopeptins that form pores (labeled with a P) in the host plasma membrane. Overall, the activities of these virulence factors may promote pathogen growth through suppression of host defenses, modulation of host cell physiology, and release of nutrients and water into the apoplast. Additionally, they may contribute to disease symptom development such as chlorosis and necrosis, thus facilitating pathogen release from infected tissue

Virulence Strategies of Plant Pathogenic Bacteria

production, motility, and virulence of Pseudomonas syringae pv. syringae (Quin˜ones et al. 2005; Dulla and Lindow 2009). Quorum sensing signaling also controls virulence determinants in soft-rot Erwinia species including production of extracellular enzymes and exopolysaccharides (EPS; discussed below), tolerance to free oxygen radicals, and subsequent symptom development in the host plant (Molina et al. 2005; Barnard and Salmond 2007). The regulatory system Phc (phenotype conversion) controls virulence factors of Ralstonia solanacearum in a populationdependent manner in the transition from soil to parasitic lifestyle (reviewed by von Bodman et al. 2003). Interestingly, the types of traits that are controlled by QS signaling are different in various bacterial species and seem to be correlated with their lifestyle.

Extracellular Polysaccharides Many plant pathogens produce large amounts of exopolysaccharides (EPSs). EPSs are carbohydrate polymers that are secreted by bacteria and form either a closely attached capsule layer surrounding the bacterial cell or a loosely associated extracellular slime (Denny 1995). The virulence of several phytopathogenic bacteria, including R. solanacearum, E. amylovora, X. campestris, and P. syringae, is associated with their ability to produce various EPS polymers during growth in plant tissue (Denny 1995). EPSs are believed to provide a selective advantage to phytopathogenic bacteria through multiple functions including (1) facilitating absorption of water, minerals and nutrients; (2) providing protection from abiotic stresses encountered during epiphytic or saprophytic growth, as well as from toxic molecules encountered during growth in plant tissue; (3) promoting colonization and spread within host tissue; and (4) contributing to the production of disease symptoms such as watersoaking and wilting (Denny 1995). One of the most important virulence-associated characteristics of the wilt pathogen R. solanacearum is the ability to produce large amounts of a viscous, high-molecular-mass, acidic EPS (EPS1) in planta. Production of large amounts of EPS1 by bacteria colonizing vascular tissue appears to interfere with transduction of water and nutrients within infected plants, resulting in wilting and, in some cases, the ultimate death of aerial portions of the plant (Denny and Baek 1991; Kao et al. 1992). Consistent with these observations, infection with R. solanacearum strains bearing mutations in the EPS1 biosynthetic loci resulted in reduced wilting (Denny and Baek 1991). A study involving detailed microscopic analysis of the infection process revealed that EPS1-deficient mutants of R. solanacearum are less invasive than wild-type strains, suggesting that EPS1 may also be required for efficient colonization and movement within plant roots (Saile et al. 1997; Araud-Razou et al. 1998). Further, the accumulation of electron-dense material in plant tissue infected with eps1 mutants raises the possibility that these mutants elicit nonspecific defenses within the host. Thus, EPS1 may also contribute to pathogen virulence by evading or suppressing host defenses (Araud-Razou et al. 1998).

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Erwinia amylovora, well known as the causal agent of fire blight of pear, produces two major EPSs, levan and amylovoran, that may contribute to this pathogen’s ability to also cause wilting diseases on young plants (Denny 1995). However, only amylovoran, a viscous, acidic heteropolysaccharide containing primarily galactose and glucuronic acid (Eastgate 2000), has been clearly demonstrated to contribute to virulence of E. amylovora, and amylovoran-negative mutants exhibit reduced in planta bacterial growth and symptom development (Bellemann and Geider 1992; Bernhard et al. 1993). Amylovoran is proposed to promote virulence by suppressing pathogen recognition by the host (Metzger et al. 1994), promoting tissue invasion and causing water-soaking and tissue collapse (Eastgate 2000). Xanthomonas campestris strains produce large amounts of the EPS known as xanthan gum that can accumulate to very high levels in infected plant tissues (Denny 1995). Xanthan gum is a high-molecular-weight EPS composed of a cellulose backbone to which trisaccharide side chains are attached. Xanthan exhibits several unique properties in solution that have rendered it useful in industrial applications (Becker et al. 1998). However, despite being one of the best studied polysaccharides produced by phytopathogenic bacteria, the role of xanthan in pathogenesis is not understood. Xanthan clearly contributes to pathogen aggressiveness, as X. campestris strains carrying mutations that specifically disrupt EPS production exhibit reduced virulence (Katzen et al. 1998). It has been proposed that xanthan contributes to X. campestris fitness by providing protection against desiccation and hydrophobic molecules and through facilitating tissue colonization by promoting adhesion of bacteria to biological surfaces (Chan and Goodwin 1999). The recent discovery that xanthan is involved in formation of aggregates of X. campestris pv. campestris in culture suggests that this EPS may be involved in biofilm formation (Dow et al. 2003). Biofilm formation may be important during early stages of tissue colonization, for example, by promoting epiphytic survival or by providing protection against antimicrobial compounds encountered within plant tissues. Interestingly, dispersal of bacteria from such a biofilm at later stages of infection may be required to facilitate colonization of the vascular system (Dow et al. 2003). The major EPS produced by P. syringae pvs. phaseolicola, lachrymans, and tomato growing in planta is alginate, a copolymer of O-acetylated b-1,4-linked D-mannuronic acid and its C-5 epimer, L-glucuronic acid (Osman et al. 1986; Fett and Dunn 1989). Studies have associated P. syringae virulence with the amount of alginate produced in culture (Osman et al. 1986; Denny 1995). A P. syringae pv. syringae alginate lyase (algL) mutant impaired in alginate production exhibited reduced epiphytic fitness, grew to lower levels in plant tissue, and elicited reduced disease symptoms on bean leaves (Yu et al. 1999). However, lack of alginate synthesis by P. syringae pv. glycinea could not be associated with reduction of in planta bacterial multiplication (Schenk et al. 2008). These findings indicate that production of alginate and its role in increased

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epiphytic fitness and pathogen virulence may be pathovar specific. Although the role of alginate in promoting virulence of P. syringae is not fully understood, alginate contributes to the virulence of the human pathogen P. aeruginosa by forming biofilms and providing protection from host defenses and antibiotic treatment (Boyd and Chakrabarty 1995; Franklin et al. 2011).

Plant Cell Wall Degrading Enzymes Plant cell walls also play an important role in plant-pathogen interactions. Unlike animal cells, plant cells are surrounded by a semirigid cell wall that provides structural support, maintains cell shape, cements adjacent plant cells together, and serves as a barrier to pathogen invasion and spread within infected tissue. Plant cell walls are composed of several complex carbohydrate polymers, the most abundant of which are cellulose, hemicelluloses, and pectin (Carpita and McCann 2000; Pauly and Keegstra 2010). As extracellular pathogens, phytopathogenic bacteria encounter plant cell walls as barriers preventing access to the cytoplasmic contents of host cells, as deterrents to pathogen spread within infected tissue, as physical substrates on which to grow, and as a potentially rich source of carbon (Agrios 2005). Thus, not surprisingly, many plant pathogens include a battery of cell wall degrading enzymes in their repertoire of virulence factors. These enzymes include pectinases (e.g., polygalacturonases, pectate lyases, and pectin methyl esterases), cellulases, and proteases that work collectively to soften or break down plant cell walls, thereby facilitating pathogen entry and the release of nutrients for pathogen growth (Barras et al. 1994). The secretion of these exoenzymes may also result in the loosening of the middle lamellae that hold together adjacent plant cell walls, thus promoting the spread of pathogens between host cells and beyond the initial infection site. The soft-rot pathogens, such as E. chrysanthemi and E. carotovora, which make their living by macerating the plants’ tissue, secrete multiple cell wall degrading enzymes. The importance of these enzymes in the virulence of these pathogens is well established (Toth et al. 2003). X. campestris pv. campestris also has an extensive collection of genes encoding putative cell wall degrading enzymes, including several pectic enzymes and cellulases (da Silva et al. 2002). Presumably, these enzymes contribute to the massive degeneration of plant tissue that occurs during development of black rot disease in plants infected with X. campestris pv. campestris (Agrios 2005). However, as Erwinia and X. campestris pathogens secrete complex mixtures of degradative enzymes and possess multiple genes encoding functionally redundant isoenzymes, the precise role of any one of these enzymes in pathogenesis has been difficult to determine (Chan and Goodwin 1999; Toth et al. 2003). The roles of plant cell wall degrading enzymes during pathogenesis of vascular wilt and leaf spotting pathogens such as R. solanacearum and P. syringae are less clear. Ralstonia solanacearum encodes multiple known or predicted pectolytic

enzymes, including endoglucanases, polygalacturonases, and a pectin methyl esterase (Genin and Boucher 2002; Salanoubat et al. 2002). Genetic studies have revealed that several of these pectolytic enzymes contribute quantitatively to bacterial wilt disease development by facilitating invasion, colonization, and systemic spread of the pathogen within host tissue (Schell et al. 1988; Huang and Allen 1997, 2000). Recent sequence analysis has revealed that P. syringae pv. tomato strain DC3000 also encodes several potential cell wall degrading enzymes, including a polygalacturonase, a pectin lyase, and three enzymes predicted to have cellulolytic activity (Buell et al. 2003). The role of these enzymes in DC3000 virulence is not known, and no cell wall degrading activity has been reported for this strain. However, pectolytic enzymes have been reported to contribute to symptom development during infection by P. syringae pv. lachrymans (Bauer and Collmer 1997). Interestingly, three TTSS effector proteins (HopPmaHPto, HrpW, and HopPtoP), classified as ‘‘helper proteins’’ that may assist in delivery of TTSS-secreted proteins, possess carboxyterminal domains with similarity to pectolytic enzymes (Charkowski et al. 1998; Boch et al. 2002; Collmer et al. 2002). The secretion of these potential pectolytic enzymes (either through sec-dependent or TTSS-dependent processes) could possibly facilitate the assembly of functional type III secretion complexes at the bacteria-plant cell wall interface.

Production of Low-Molecular-Weight Phytotoxins Many plant pathogens produce low-molecular-weight, nonhost specific phytotoxins that are not essential for pathogenicity but yet contribute to bacterial virulence and increase disease symptoms such as chlorosis and necrosis. These toxins act either by directly damaging plant cells or by modulating host cellular metabolism to promote symptom development (Alfano and Collmer 1996; Bender et al. 1999; Birch 2001). The best characterized of these phytotoxins are those produced by P. syringae species and include lipodepsipeptide toxins (e.g., syringomycins and syringopeptins), modified peptides (e.g., tabtoxin and phaseolotoxin), polyketides (e.g., coronatine), and a combination of peptides and polyketides (e.g., syringolin group).

Lipodepsipeptide Toxins Syringomycins and syringopeptins are examples of the two classes of lipodepsipeptide toxins produced by P. syringae pv. syringae during infection. They are synthesized by nonribosomal peptide synthetases which are encoded by the syr and syp genomic islands of P. syringae pv. syringae. The genomic structure of these genomic islands including the regulatory network controlling toxin production has been determined (Scholz-Schroeder et al. 2003; Lu et al. 2005). The syringomycins are cyclic lipodepsinonapeptide phytotoxins that consist of a polar cyclic

Virulence Strategies of Plant Pathogenic Bacteria

peptide head containing nine amino acids attached to a hydrophobic 3-hydroxy carboxylic acid tail (Bender and Scholz-Schroeder 2004). Several structurally similar syringomycins are produced by different P. syringae pv. syringae strains that contain different amino acid residues in the nine-peptide ring. Syringopeptins are larger than the syringomycins and contain a peptide moiety of 22 or 25 amino acids attached to either a 3-hydroxydecanoic or a 3-hydroxydodecanoic acid (Bender and Scholz-Schroeder 2004). As in syringomycins, the amino acid chain is cyclized to form a nine-peptide ring. Many of the amino acids present in the syringopeptins are hydrophobic and thus contribute to the amphipathic nature of these toxins. The syringomycins and syringopeptins induce necrosis in plant tissues by forming pores in the plant cell plasma membrane possibly through a mechanism involving initial insertion of toxin monomers into the membrane, followed by aggregation of multiple monomers to form a pore (Bender and ScholzSchroeder 2004). The formation of pores in lipid membranes increases transmembrane ion flux, causing disruption of membrane electrical potential and eventual plant cell death (Hutchison and Gross 1997). The amphipathic nature of these toxins is likely to facilitate their insertion into plant cell membranes. Lipodepsipeptide phytotoxins are likely to play an important role in interactions between P. syringae pv. syringae and its hosts, as all strains of P. syringae pv. syringae analyzed to date produce both syringomycins and syringopeptins. Genes encoding these toxins are induced by plant-derived phenolic compounds (Wang et al. 2006), and they contribute quantitatively to P. syringae pv. syringae virulence; however, the relative importance of these toxins vary among different pathogen-host interactions (Bender et al. 1999; Scholz-Schroeder et al. 2001). Much progress has been made toward understanding the biosynthesis and pore-forming activities of the lipodepsipeptide phytotoxins (Bender and Scholz-Schroeder 2004), and they contribute to pathogen virulence by stimulating plant cell necrosis and disease lesion development. Additionally, as both syringomycins and syringopeptins exhibit biosurfactant activities, they could potentially contribute to virulence by reducing the surface tension of water and thus facilitate the spread of bacteria across plant surfaces (Bender et al. 1999), thereby promoting tissue colonization and spread of the pathogen within infected plant tissue. Interestingly, the recent completion of the genomic sequence of R. solanacearum has revealed two large open reading frames predicted to encode proteins with high similarity to syringomycin synthase (Salanoubat et al. 2002), suggesting that this pathogen may also produce syringomycin.

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phaseolotoxin produced by P. syringae pvs. phaseolicola and actinidae (Mitchell 1976; Bender et al. 1999), tabtoxin produced by P. syringae pvs. tabaci, coronafaciens, and garcae (Uchytil and Durbin 1980; Moore et al. 1984), and mangotoxin produced by P. syringae pvs. syringae and avellanae (Gasson 1980; Arrebola et al. 2003). These toxins can interfere with host nitrogen metabolism by inhibiting specific enzymes involved in the biosynthesis of essential amino acids, leading to amino acid deficiencies and concomitant accumulation of nitrogen-containing intermediates that can be utilized by the pathogens (Snoeijers et al. 2000; Arrebola et al. 2011). Tabtoxin is a dipeptide toxin that contains tabtoxinine-blactam (TbL), linked by a peptide bond to threonine (Bender et al. 1999). TbL is the toxic moiety of tabtoxin and is released from the intact toxin upon hydrolysis of the peptide bond by the action of aminopeptidases within the plant (Levi and Durbin 1986). TbL, which induces the degradation of chlorophyll in plant cells (thus causing yellowing or ‘‘chlorosis’’), irreversibly inhibits the enzyme glutamine synthetase (Thomas et al. 1983). As glutamine synthetase is required for efficient detoxification of ammonia in plant cells, inactivation of this enzyme results in accumulation of high levels of ammonia and the disruption of thylakoid membranes within the chloroplast. Phaseolotoxin is a tripeptide consisting of ornithine, alanine, and a homoarginine linked to a sulfo-diaminophosphinyl moiety (Moore et al. 1984). When taken up by plant cells, phaseolotoxin is hydrolyzed to produce octicidine, an irreversible inhibitor of ornithine carbamoyl transferase (OCTase; Mitchell and Bieleski 1977). OCTase is a key enzyme in the urea cycle that converts ornithine and carbamoyl phosphate to citrulline. Inhibition of OCTase by phaseolotoxin results in accumulation of ornithine and reduction in arginine levels, leading to the production of severe chlorosis within plant tissue (Bender et al. 1999). Both tabtoxin and phaseolotoxin contribute significantly to pathogen virulence, presumably by inhibiting photosynthesis and thus limiting available resources within the plant for mounting a successful defense response and by contributing to the severe yellowing of plant tissues associated with disease (Agrios 2005). The structure of mangotoxin has not been conclusively determined yet; however, it is most likely to consist of two amino acids linked by a sugar residue (Arrebola et al. 2003). It inhibits the ornithine N-acetyltransferase (OAT) that converts a-N-acetyl-ornithine to L-ornithine during arginine biosynthesis. The gene cluster encoding enzymes for mangotoxin biosynthesis has been identified (Arrebola et al. 2011). Studies with the mgoA mutant of P. syringae pv. syringae reveal that it is incapable of producing mangotoxin and shows reduced virulence on tomato leaves (Arrebola et al. 2007).

Modified Peptide Toxins with Antimetabolite Activity

Coronatine

Antimetabolite toxins are oligopeptides produced by several pathovars of P. syringae that can inhibit host metabolite pathways. The most well-studied antimetabolite phytotoxins include

Coronatine is produced by several P. syringae strains (Bender et al. 1999) and contributes to the virulence of P. syringae via several mechanisms. It plays important roles in early and late

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stages of disease development including suppression of stomatal immunity and thus enhancing entry into host tissue (Melotto et al. 2006; Zeng et al. 2010); promoting pathogen growth in the apoplast subsequent to entry (Zeng et al. 2010); and enhancing disease symptom development (Brooks et al. 2005; Melotto et al. 2008a). Results from genetic studies, utilizing both P. syringae mutants impaired in coronatine biosynthesis and Arabidopsis and tomato mutants impaired in JA signaling, indicate that coronatine promotes pathogen virulence by stimulating JA signaling within the plant (Feys et al. 1994; Kloek et al. 2001; Zhao et al. 2003; Brooks et al. 2005; Laurie-Berry et al. 2006). These studies revealed that SA- and JA-dependent defense signaling pathways are mutually antagonistic; coronatine-induced activation of JA signaling results in inhibition of SA-dependent defense responses, which are effective in limiting P. syringae infection and disease (Brooks et al. 2005; Laurie-Berry et al. 2006; Uppalapati et al. 2007). In support of this antagonist interaction between JA and SA signaling, reduced susceptibility to P. syringae in Arabidopsis coronatine insensitive1 (coi1) and JA insensitive1 (jin1/myc2) mutant plants is associated with increased signaling through the SA-dependent defense pathway (Kloek et al. 2001; Nickstadt et al. 2004; Laurie-Berry et al. 2006), and coronatine suppresses induction of several SA-dependent defense-related genes in tomato (Zhao et al. 2003; Uppalapati et al. 2005). Thus, coronatine suppresses SA-mediated host defenses, thereby providing P. syringae with a window of opportunity during which it can colonize and grow within host tissue. Coronatine may also be directly involved in disease symptom development via multiple SA-independent processes. The in planta growth defect of P. syringae coronatine biosynthetic mutants is suppressed in Arabidopsis mutants in which SA-dependent defenses are compromised (Brooks et al. 2005; Zeng et al. 2010). However, although the coronatine mutants grow to wild-type levels in SA-deficient plants, disease symptom development is not fully restored (Brooks et al. 2005). Recent studies suggest that coronatine may contribute to disease symptom production via two mechanisms: (1) by stimulating production of reactive oxygen species that lead to disease-associated necrotic cell death (Ishiga et al. 2009) and (2) by stimulating expression of the STAYGREEN (SGR) gene, which results in enhanced chlorophyll degradation and chlorosis of the infected tissue (Mecey et al. 2011). At the molecular level, the polyketide phytotoxin coronatine is of interest to both plant biologists and plant pathologists, as it is a structural and functional mimic of the endogenous plant hormone jasmonyl-L-isoleucine (JA-Ile) (Staswick and Tiryaki 2004; Fonseca et al. 2009). Coronatine is comprised of two distinct chemical moieties, coronafacic acid (CFA; a polyketide) that shares structural and functional relatedness with jasmonic acid (JA) and coronamic acid (CMA; an ethylcyclopropyl amino acid) that is a cyclized form of isoleucine. CFA and CMA are joined by an amide linkage forming the intact coronatine molecule (Bender et al. 1999), which is required for full virulence of P. syringae (Brooks et al. 2005; Uppalapati et al. 2007). Specifically, the intact coronatine

molecule closely resembles the active form of the plant jasmonate (+)-7-iso-JA–L-Ile (Fonseca et al. 2009). JA-Ile is a member of a family of fatty acid-derived signaling molecules, the jasmonates, which play an important role in many aspects of plant growth, development, and defense (Wasternack 2007; Browse 2009). The biological effects of coronatine closely resemble those induced by jasmonates and include induction of chlorosis, production of the protective pigment anthocyanin, inhibition of root growth, promotion of plant cell growth, and the induction of several JA-responsive genes (Feys et al. 1994; Bender et al. 1999; Zhao et al. 2003; Uppalapati et al. 2005). The most compelling evidence that coronatine is a functional analog of JA-Ile is the observation that it competitively binds to the JA-Ile receptor complex. This receptor complex is formed by the CORONATINE INSENSITIVE1 (COI1) and JASMONATE ZIM-DOMAIN (JAZ) proteins (Katsir et al. 2008; Yan et al. 2009; Melotto et al. 2008b; Sheard et al. 2010). COI1 is the F-box protein in the SCFCOI1 ubiquitin E3 ligase complex that targets JAZ proteins for degradation (Chini et al. 2007; Thines et al. 2007). Binding of JAZ proteins to the SCFCOI1 complex is stabilized in the presence of coronatine (or JA-Ile), leading to the degradation of JAZ proteins, which are transcriptional repressors of JA signaling. JAZ protein degradation, therefore, results in the expression of JA-responsive genes and the activation of JA-mediated responses leading to disease progression through a yet-to-bediscovered mechanism(s).

Syringolin Group Phytotoxins in the syringolin group contain a 12-membered ring structure consisting of 5-methyl-4-amino-2-hexenoic acid, 3, 4-dehydrolysine, and a dipeptide tail. The substitution of one of more of these amino acids forms syringolin A–F (Wa¨spi et al. 1998). Syringolin A is the best characterized among this group of small cyclic tripeptide phytotoxins produced by P. syringae pv. syringae. Recently, it has been shown that syringolin A negatively acts on the catalytic activity of eukaryotic proteasomes (Groll et al. 2008) and functions in disease development. Surface inoculation of a syringolin A-deficient strain of P. syringae pv. syringae B728a shows reduced bacterial multiplication and symptoms on the host plant Phaseolus vulgaris (Groll et al. 2008; Schellenberg et al. 2010). Similar to coronatine, syringolin A also stimulates opening of the stomatal pore and belongs to a growing group of antistomate defense factors (Melotto et al. 2006; Gudesblat et al. 2009; Schellenberg et al. 2010). Syringolin A-deficient bacteria are unable to open stomata of bean leaves; however, this effect can be reverted by exogenous application of syringolin A or the proteasome inhibitor MG132, suggesting that syringolin A acts through protein turnover affecting guard cell movement (Schellenberg et al. 2010). Interestingly, the defense hormone SA and the SA signaling component NPR1 (non-expressor of pathogenesis-related genes) are required for stomatal immunity (Melotto et al. 2006; Zeng and He 2010).

Virulence Strategies of Plant Pathogenic Bacteria

NPR1 is a transcription activator of systemic acquired resistance that is regulated in part through protein turnover, and it is degraded through the proteasome (Spoel et al. 2009; Trujillo and Shirasu 2010) suggesting that several pathogen-produced phytotoxins act as antistomate defense factors, at least in part, by blocking NPR1-dependent SA signaling.

Type III Secretion Although bacterial pathogens of plants such as P. syringae and Xanthomonas have multiple secretion pathways as revealed by genome analysis (Buell et al. 2003; Cunnac et al. 2009; Ryan et al. 2011), like most gram-negative bacterial pathogens of animals, the bacterial plant pathogens discussed in this chapter require a functional type III secretion system (TTSS) for pathogenesis (Galan and Collmer 1999; Cornelis and Van Gijsegem 2000; Staskawicz et al. 2001; Buttner and Bonas 2003; Jin et al. 2003a; Cunnac et al. 2009; Ryan et al. 2011; Lindeberg et al. 2012; Buttner and He 2009). Type III secretion systems mediate the transfer of bacterial proteins (also referred to as ‘‘effectors’’) directly into the cytosol of the host cell, where they interfere with or modulate normal host cell processes to facilitate bacterial invasion, growth, and disease production (> Fig. 2.1). Plant pathogenic bacterial mutants defective in TTSS are usually unable to grow or cause disease on normally susceptible hosts, indicating that the integrity of the TTSS is essential for pathogenesis (Mudgett 2005; Boch and Bonas 2010; Lindeberg et al. 2012). Much progress has been made toward elucidating the structure and components of the TTSS and in identifying the effector proteins secreted through this apparatus. Insights into the function of several type III effectors and how they contribute to the virulence of plant pathogens have also been recently obtained.

Structure and Components of TTSS of Bacterial Plant Pathogens The structural components of the TTSS of gram-negative bacterial pathogens of animal and plants are highly conserved. These systems have been extensively described elsewhere (Galan and Collmer 1999; Collmer et al. 2000; Cornelis and Van Gijsegem 2000; Buttner and Bonas 2002; Jin et al. 2003a; Buttner and He 2009); therefore, the TTSS apparatus will not be discussed in detail. However, note that structurally the TTSSs of plant pathogenic bacteria are slightly different from those described for the animal pathogens. For example, the TTSS of several mammalian pathogens, such as Salmonella enterica serovar Typhimurium and Shigella flexneri, are associated with protruding, needle-like surface structures that are approximately 80 nm in length (Kubori et al. 1998; Blocker et al. 1999). The TTSS of several plant pathogenic bacteria are associated with relatively longer, pilus-like structures referred to as ‘‘Hrp pili’’ (He and Jin 2003). The Hrp pilus of P. syringae pv. tomato strain DC3000 is approximately 8 nm in diameter and has been observed to be up to 200 nm in length, which is

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presumably long enough to span the plant cell wall (Brown et al. 2001; Jin and He 2001). Several studies suggest (but do not directly demonstrate) that the Hrp pilus serves as the conduit through which bacterial proteins are secreted (Brown et al. 2001; Jin and He 2001).

Identification of Type III Effectors The importance of TTSS for pathogenesis has prompted many research groups to direct a significant amount of effort toward identifying and characterizing proteins that are secreted through the TTSS. An inventory of effector proteins secreted by plant pathogenic bacteria has been compiled in several excellent reviews (Collmer et al. 2002; Buttner and Bonas 2003; Buttner et al. 2003; Greenberg and Vinatzer 2003; Jin et al. 2003a, Cunnac et al. 2009; Lindeberg et al. 2012). A variety of approaches have been used to identify these effectors. The first type III-secreted proteins studied were those identified on the basis of their ability to elicit TTSS-dependent host defense responses on resistant plant genotypes (Staskawicz 2001). This may not be surprising, given the eagerness of plant pathologists to elucidate the mechanisms underlying pathogen recognition and disease resistance. Further, given that type III effectors are secreted directly into host cells, and thus may serve as ‘‘easy targets’’ for recognition during the evolution of host surveillance systems, it may not be surprising that many effector molecules serve as elicitors of plant defense. Recently, more comprehensive approaches for identifying genes encoding type III effectors have been employed. These approaches include (1) utilizing information regarding gene location and gene regulation, (2) direct functional assays to screen for secreted proteins, and (3) taking advantage of common structural features of known TTSS-secreted proteins to carry out ‘‘genomic mining’’ experiments. For instance, in P. syringae and X. campestris, several genes encoding effector proteins are located within or adjacent to the gene clusters encoding the structural components of the TTSS (Alfano et al. 2000; Noel et al. 2002; Charity et al. 2003). Further, the expression of many P. syringae genes encoding either structural TTSS components or TTSS effector proteins depends on HrpL, an alternative RNA polymerase s factor required for pathogenesis (Xiao et al. 1994). The HrpL s factor directs transcription of TTSS-associated genes by recognizing a consensus ‘‘hrp box’’ in the promoter regions of these genes (Innes et al. 1993; Xiao and Hutcheson 1994). This information has been used as the basis of genetic screens to identify genes potentially encoding TTSS effector proteins (Fouts et al. 2002; Zwiesler-Vollick et al. 2002; Bretz et al. 2003). Likewise, a molecular genetic strategy has been used to identify X. campestris genes whose expression is dependent on HrpX, an AraC-like transcriptional activator that is essential for induction of TTSS-related genes in this organism (Noel et al. 2001). Although type III effectors do not have an obvious signal sequence targeting them for secretion, the proteins are modular

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in structure, with the amino terminal region carrying information required for secretion (Guttman et al. 2002). Analyses of the primary amino acid sequences of several known P. syringae type III effectors has revealed a strikingly well-conserved pattern of amino acid biases within the first 50 residues that is essential for secretion (Guttman et al. 2002; Petnicki-Ocwieja et al. 2002). As the genomes of several plant pathogenic bacteria have been sequenced, the above features have facilitated genomic mining experiments to identify the entire repertoire of type III effectors secreted by these pathogens (Collmer et al. 2002; Greenberg and Vinatzer 2003; Ryan et al. 2011; Potnis et al. 2011; Lindeberg et al. 2012). Interestingly, the available genomes of plant pathogenic bacteria revealed large inventories of type III effectors. For instance, P. syringae pv. tomato strain DC3000 and pv. phaseolicola race 6 have 29 and 19 effector proteins, respectively, that are translocated to and elicit hypersensitive response in plant cells as detected by a variety of functional screens (Chang et al. 2005; Cunnac et al. 2009) The fact that plant pathogenic bacteria secrete many type III effectors could be an adaptive feature of plant pathogens and suggests that functional redundancy may exist among these effectors (Buttner and Bonas 2003). Consistent with this hypothesis, mutations in single effector genes usually do not dramatically alter bacterial virulence, at least when assayed under laboratory conditions (Ponciano et al. 2003). In contrast to TTSS structural components, which are highly conserved between various plants and animals, the sequences and inventories of type III effectors vary considerably among different plant pathogens and even among different strains of the same species (Collmer et al. 2002; Greenberg and Vinatzer 2003). This variation suggests that different strains have evolved different repertoires of virulence factors to infect and cause disease on specific host plants. Thus, characterizing type III effectors and elucidating the mechanisms through which they contribute to pathogenesis is of great interest and may eventually provide insight into the molecular basis of host specificity.

Elucidating the Function of Type III Effectors The majority of TTSS effectors secreted by bacterial plant pathogens are predicted to function inside plant cells, and secretion into the host cell has been experimentally demonstrated for many effector proteins (Casper-Lindley et al. 2002; Szurek et al. 2002; Hotson et al. 2003; Chang et al. 2005; Cunnac et al. 2009). Several type III effectors have been directly demonstrated to have elicitor and virulence activities inside plant cells (Leister et al. 1996; Chen et al. 2000; Marois et al. 2002; Hauck et al. 2003; Jamir et al. 2004; Block and Alfano 2011), and are predicted to modulate various aspects of host cell biology and physiology to support bacterial virulence, proliferation, dissemination, and promote disease (Alfano and Collmer 1996; Greenberg and Vinatzer 2003; Jin et al. 2003a; Ponciano et al. 2003; Boch and Bonas 2010; Block and Alfano 2011). Various strategies to elucidate the activities of these effectors have been employed, including protein sequence and structural

analyses, biochemical approaches to identify interacting proteins, and the analysis of transgenic plants expressing effector proteins. Such studies have revealed that, despite their prokaryotic origin, many type III effectors have features typical of eukaryotic proteins, consistent with their activity within plant cells. For instance, the P. syringae effector proteins AvrRpm1, AvrB, AvrPto, and AvrPphB have consensus N-terminal myristoylation sites that are myristoylated inside host cells (Nimchuk et al. 2000). This modification is required for the proper localization of these type III effectors at the host plasma membrane (Nimchuk et al. 2000; Shan et al. 2000). Another effector from P. syringae, HopI1, is targeted to the chloroplast and suppresses the accumulation of the defense hormone SA (Jelenska et al. 2007). Furthermore, all members of the AvrBs3/ PthA family of effectors and XopD, found in the genus Xanthomonas, carry a functional nuclear localization signal (NLS), share structural features of eukaryotic transcription factors, and are classified as the ‘‘transcription activator-like’’ (TAL) family (Schornack et al. 2006; Kim et al. 2008). An NLS at the carboxy-terminus of AvrBs3 from X. campestris is required for interaction with importin a, which is part of the host nuclear import machinery (Szurek et al. 2001). The function of these Xanthomonas effectors, several of which activate host gene expression, either to promote virulence or to trigger defense responses, have recently been comprehensively reviewed (Boch and Bonas 2010; Rivas and Genin 2011; Ryan et al. 2011). These studies illustrate that type III effectors of prokaryotic origin have evolved to take advantage of eukaryote-specific post-translational modification and targeting mechanisms to access specific subcellular compartments within the host cell. As is described in more detail below, the molecular activities identified to date for TSS effectors include facilitating type III secretion; suppressing plant immune defense by altering host cell biology at various levels; causing proteolysis of host proteins; regulating host transcription; modulating plant RNA metabolism and vesicle trafficking; and altering plant hormone synthesis, homeostasis, and signaling. Facilitators of Type III Secretion. Several type III effector proteins, known as hairpins, are secreted into the apoplast and function as ‘‘helper proteins’’ to facilitate translocation of type III secretion effectors through the host plasma membrane during pathogenesis. Some examples are HrpZ and HrpW of P. syringae (Lee et al. 2001), HrpF of X. campestris pv. vesicatoria (Buttner et al. 2002), and HrpN of E. amylovora (Bocsanczy et al. 2008). The P. syringae HrpZ protein forms oligomers and has strong affinity for the plasma membrane lipid phosphatidic acid (Haapalainen et al. 2011). Likewise, HrpF from X. campestris contains two putative transmembrane regions, suggesting its association with membranes (Buttner et al. 2002), and is required to translocate type III effectors essential for pathogenicity in the host plant (Jiang et al. 2009). Both proteins have lipid-binding activity and form ion-conducting pores in vitro when associated with lipid bilayers (Lee et al. 2001; Buttner et al. 2002). The pore-forming activity of these proteins suggests that they function in assisting delivery of effectors into the plant cell cytoplasm and/or by mediating nutrient and water release from

Virulence Strategies of Plant Pathogenic Bacteria

host cells. Since HrpF is dispensable for protein secretion in vitro, but is required for the recognition of an effector with elicitor activity in vivo, it has been proposed that HrpF may facilitate translocation of one or more effector proteins into the host cell (Rossier et al. 2000). Suppression of Host Immunity. Plants possess a complex immune system that efficiently detects potential microbial invaders and fends off dangerous microbes. Plant innate immunity is associated with perception of pathogen- or microbeassociated molecular patterns (PAMPs or MAMPs) by pattern recognition receptors (PRRs) at the plant’s cell surface. For example, two well-characterized MAMPs, the flagellin peptide flg22 and elongation factor Tu (EF-Tu), are detected by the PRRs FLS2 and EFR1, respectively (Zipfel et al. 2004, 2006). This branch of innate immunity is also known as PTI (PAMPtriggered immunity; Jones and Dangl 2006; Boller and Felix 2009). Several type III effectors can effectively suppress PTI, and this is hypothesized to be fundamental in the evolution of pathogen virulence (Jones and Dangl 2006). However, pathogens do not always cause disease, as plants have evolved another level of innate immunity known as ETI (effector-triggered immunity), formerly referred to as ‘‘gene-for-gene resistance’’ (Jones and Dangl 2006; Thomma et al. 2011). ETI is established by either direct binding of host resistance proteins to bacterial effectors or by host resistance proteins monitoring the modification of (or ‘‘guarding’’) effector targets in the host cell. Although the latter mechanism appears to be more common than the former, in both cases, a rapid, localized host cell death, known as the hypersensitive responses (HR), is typical of ETI (Lewis et al. 2009). Plant host immunity and the molecular activities of bacterial effectors suppressing both PTI and ETI are the subject of several excellent, recent reviews (Chisholm et al. 2006; Jones and Dangl 2006; Block et al. 2008; Boller and He 2009; Lewis et al. 2009; Block and Alfano 2011; Chen and Ronalds 2011; Spoel and Dong 2012). Proteolysis of Host Proteins. Several type III effectors have proteolytic activity that interferes with normal host physiology and actively suppresses plant immunity. For instance, two effectors from P. syringae pv. tomato strain DC3000, HopM1 and AvrPtoB, promote degradation of host proteins through the 26S proteasome pathway. HopM1 interferes with vesicle trafficking of possible defense cargoes to the plasma membrane by triggering the degradation of the ARF guanine exchange factor AtMIN7 through the proteasome pathway. Plants lacking AtMIN7 are exceedingly susceptible to bacteria (Nomura et al. 2006). The specificity of ubiquitinylation of proteins targeted for degradation in eukaryotes is controlled by a variety of E3 ligase complexes. Of relevance to this discussion is the SCF-type E3 ubiquitin ligase complex, containing Skp1, Cullin1, and an F-box protein subunit. Interestingly, some bacterial effectors seem to highjack the host SCF-type E3 ubiquitin ligase mechanism to promote disease. For instance, AvrPtoB has an E3 ligase activity that targets the PAMP receptors FLS2, ERF1, CERK1, and the receptor partner BAK1 thus suppressing plant immunity in vivo (Gohre et al. 2008; Shan et al. 2008; Gimenez-Ibanez et al. 2009). The GALA effectors from R. solanacearum contain an

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F-box domain that is required for several stages of disease development on plants (Angot et al. 2006; Turner et al. 2009). The host targets of GALA effectors remain to be identified. The AvrPphB cysteine protease from P. syringae pv. phaseolicola cleaves both itself and PBS1, an Arabidopsis protein kinase required for ETI (Shao et al. 2003). Interestingly, AvrPphB can inhibit PTI by cleaving PBS1-like kinases (Zhang et al. 2010) that interact with FLS2, demonstrating that PTI and ETI are connected by the activities of pathogen effector proteins. A model for the action of AvrPphB is still under development and has been recently reviewed by Day and He (2010). AvrRpt2 from P. syringae pv. tomato is a cysteine protease that cleaves itself to form a mature protease inside the host plant (Jin et al. 2003b) as well as several host proteins, including the Arabidopsis RIN4 protein, a negative regulator of PTI and ETI (Axtell and Staskawicz 2003; Axtell et al. 2003; Chisholm et al. 2005; Day et al. 2005; Kim et al. 2005; Dodds and Rathjen 2010). As will be discussed further below, AvrRpt2 also promotes pathogen virulence and modulates auxin sensitivity in the host. However, it is not clear whether the cysteine protease activity of AvrRpt2 is also required for both of these activities. RIN4 appears to be an important virulence target or defense decoy, as it is also targeted by several other type III effectors (AvrB, AvrRpm1, and HopF2). However, it is not clear how alteration of RIN4 protein enhances bacterial virulence (Dodds and Rathjen 2010). The C-terminal portion of the XopD protein from X. campestris pv. vesicatoria has a high degree of similarity with the C-terminal catalytic domain of the Ulp1 ubiquitin-like protease protein family and has a cysteine protease activity specific for small ubiquitin-like modifier (SUMO)-lated substrates found specifically in plants (Hotson et al. 2003). The XopD SUMO protease effector promotes pathogen growth and delays the onset of symptoms in the host plant tomato (Kim et al. 2008). On the basis of amino acid sequence similarity, three additional effectors from X. campestris pv. vesicatoria, AvrRxv, AvrBsT, and AvrXv4, as well as PopP1 from R. solanacearum appear to belong to the YopJ family of ubiquitin-like protein proteases. Interestingly, like XopD, YopJ exhibits specificity for SUMO-lated proteins (Orth et al. 2000; Lavie et al. 2002). Although the importance of the above demonstrated or predicted proteolytic activities in pathogen virulence in some cases is not clear, these findings suggest that the use of type III effectors to cleave specific host signaling molecules is a common strategy of bacterial pathogens. Analysis of Type III Effector Function. The findings that the primary amino acid sequences of most effector proteins do not provide much insight regarding function and that, in most cases, mutation of the effector genes does not result in pronounced virulence phenotypes have not helped to hasten our understanding of these proteins. One of the major challenges in understanding TTSS effector function is to identify the targets of these virulence factors within the host and to elucidate the roles of these molecules in pathogenesis. Despite these challenges, the scientific community has recently made tremendous progress in this area and has gained much knowledge about the function of

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TTSS effectors. Importantly, new components of the plant innate immune system, such as vesicle trafficking (Nomura et al. 2006, 2011) and RNA metabolism (Fu et al. 2007), have been identified in the search for host targets of bacterial effectors (Speth et al. 2007). Although a tremendous amount of progress toward identifying and characterizing bacterial type III effectors has been recently made, the mode of action of most of these effectors is still unknown. Future studies involving host gene expression profiling and proteomic approaches to reveal the potential effects of these pathogen molecules on plant defense and other aspects of host physiology may help elucidate the function of some of these effectors. Likewise, localization of effector proteins within host cells, as well as identification of plant proteins that interact with the effectors, will also continue to contribute to our understanding of effector function within plant cells. One experimental approach that has provided important new insight into the function of several effector proteins has been to express them ectopically in transgenic plants. Several such studies have revealed that some effectors modulate host hormone biology. For example, transgenic Arabidopsis expressing the P. syringae effector AvrRpt2 exhibit increased sensitivity to the plant hormone auxin, suggesting that AvrRpt2 modulates host auxin physiology to promote pathogen virulence (Chen et al. 2007). Likewise, expression of AvrPtoB and HopAM1 in planta results in increased levels of abscisic acid (ABA; Goel et al. 2008) and hypersensitivity to ABA, respectively (Goel et al. 2008; de Torres-Zabala et al. 2007). Thus, one role of type III effectors appears to be modulation of hormone production and/or signaling in the host. Further discussion as to how altering host hormone physiology might contribute to pathogenesis is provided below.

Modulation of Plant Hormone Homeostasis and Signaling Plant hormones, which are also often referred to as ‘‘plant growth regulators,’’ are endogenous signaling molecules important for many aspects of plant growth and development. The best known phytohormones are auxin, ethylene, cytokinins, abscisic acid (ABA), and gibberellins. More recently, several additional chemicals, including jasmonates (JA), salicylic acid (SA), brassinosteroids, nitric oxide, and strigolactones, have been recognized as phytohormones (Santner and Estelle 2009). Three of these hormones in particular, SA, JA, and ethylene, are important in mediating plant defenses in response to pathogen or herbivore attack (Hammond-Kosack and Jones 2000; Kunkel and Brooks 2002; Kazan and Manners 2009, 2012; Spoel and Dong 2009; Robert-Seilaniantz et al. 2011). Other hormones that have been implicated in plant-pathogen interactions are ABA, auxins, gibberellins, cytokinins, and brassinosteroids; their complex signaling network is nicely reviewed by Pieterse et al. (2009) and Choi et al. (2011). It is not surprising that many plant pathogens have evolved mechanisms for modulating endogenous hormone signaling

and homeostasis in their hosts, presumably as a mechanism for promoting pathogenesis. However, the specific hormone signaling pathway(s) targeted by a given pathogen, and the manner in which they are modulated seems to depend on the virulence strategy employed by the pathogen, that is, whether it is a necrotroph that rapidly kills plant cells to obtain nutrients or a biotroph that colonizes living plant tissue. Several excellent reviews on this subject are available (Spoel and Dong 2009; Grant and Jones 2009; Bari and Jones 2009; Choi et al. 2011; Robert-Seilaniantz et al. 2011; Kazan and Manners 2012). In general, there appear to be three basic strategies utilized by pathogens to manipulate the hormone physiology of their plants hosts: (1) synthesis of the hormone, or a hormone mimic; (2) perturbing hormone homeostasis (e.g., metabolism or conversion into inactive forms); and (3) modulating hormone signaling pathways. As this newly emerging paradigm has been recently discussed in several excellent reviews (Kazan and Manners 2009, 2012; Spoel and Dong 2009; Robert-Seilaniantz et al. 2011), we briefly discuss examples of these processes below.

Salicylic Acid Salicylic acid (SA) plays a central role in defense against pathogen attack. During infection, plants often accumulate SA, and exogenous application of SA or SA analogs results in enhanced resistance to a wide variety of pathogens (Durrant and Dong 2004; Bari and Jones 2009; Spoel and Dong 2009). Plant mutants that are impaired in their ability to accumulate SA exhibit enhanced susceptibility to many pathogens (Nawrath and Metraux 1999; Wildermuth et al. 2001). Thus, to successfully colonize host tissue, virulent bacterial pathogens presumably have evolved mechanisms for interfering with SA-mediated defense responses, for instance, by delaying or preventing the accumulation of high levels of SA within host tissue or by suppressing SA-dependent signaling downstream of SA accumulation. For example, the deployment of several P. syringae type III effectors, including AvrRpt2, AvrPphC, VirPphA, and AvrPphF, delays the accumulation of SA within the infected plant by inhibiting host recognition of bacteria expressing specific avirulence factors (Ritter and Dangl 1996; Jackson et al. 1999; Chen et al. 2000; Tsiamis et al. 2000; Cunnac et al. 2009; Block and Alfano 2011). Another potential strategy for interfering with induction of SA-dependent defenses is degradation of SA. This mechanism may be deployed by R. solanacearum, whose genome includes several genes encoding putative SAdegrading enzymes (Salanoubat et al. 2002). Pseudomonas syringae, and presumably other pathogens, may also facilitate colonization of host tissue by suppressing SA synthesis and signaling. As discussed above, coronatine appears to be utilized by P. syringae to downregulate SAdependent defense responses (Brooks et al. 2005; Laurie-Berry et al. 2006). Likewise, the P. syringae type III effectors AvrRpt2 and HopPtoD2 suppress the expression of SA-regulated defenserelated genes during infection on susceptible plants (Bretz et al. 2003; Chen et al. 2004). In the case of AvrRpt2, this appears to

Virulence Strategies of Plant Pathogenic Bacteria

occur without altering SA levels (Chen et al. 2004). The HopI1 type III effector acts directly at the site of SA synthesis, the chloroplast, disrupting thylakoid structure and suppressing SA accumulation (Jelenska et al. 2007). Therefore, plant pathogenic bacteria appear to deploy several different strategies to interfere with various aspects of SA-dependent defenses within the host.

Jasmonates A group of biochemically related plant growth regulators collectively referred to as ‘‘jasmonates’’ (‘‘JAs’’) are involved in defense against both herbivorous insect pests and necrotrophic bacterial and fungal pathogens that colonize dead plant tissues. Thus, intact JA signaling processes are required for resistance to attack by these organisms (Browse 2009). In contrast, JA signaling is required for disease susceptibility of Arabidopsis and tomato plants to the biotrophic bacterial pathogen P. syringae (Feys et al. 1994; Kloek et al. 2001; Zhao et al. 2003; Nickstadt et al. 2004; Laurie-Berry et al. 2006). This may not be surprising, as discussed earlier, coronatine, an important virulence factor for P. syringae, is a molecular mimic of JA-Ile (Melotto et al. 2008a; Staswick 2008; Browse 2009). Coronatine stimulates JA signaling within the plant, by binding to the receptor complex COI1/ JAZ (Katsir et al. 2008; Sheard et al. 2010). COI1-dependent activation of JA signaling is required to promote pathogenesis and disease development (Feys et al. 1994; Weiler et al. 1994; Kloek et al. 2001; Zhao et al. 2003). However, the molecular mechanism(s) underlying this process downstream of coronatine perception by the COI1/JAZ receptors is not well understood. Mounting evidence suggests that stimulation of JA signaling results in antagonism of SA-dependent defenses thereby promoting pathogen growth (Kloek et al. 2001; Zhao et al. 2003; Brooks et al. 2005; Laurie-Berry et al. 2006; Uppalapati et al. 2007; Zheng et al. 2012). Additionally, stimulation of JA appears to result in enhanced disease production, via a mechanism that is independent of SA (Brooks, et al. 2005; Laurie-Berry et al. 2006).

Ethylene The role of the gaseous plant hormone ethylene in plantmicrobe interactions is complex, as it is required for resistance against some pathogens and for disease susceptibility in others (Kunkel and Brooks 2002; Van Loon et al. 2006; RobertSeilaniantz et al. 2011). Like JA, in general, ethylene contributes to defense against necrotrophic pathogens but promotes disease in plants infected by biotrophic pathogens. Ralstonia solanacearum and P. syringae, two pathogens for which normal ethylene responsiveness in the host is important for disease development (Bent et al. 1992; Lund et al. 1998; Hoffman et al. 1999; Weingart et al. 2001; Hirsch et al. 2002), have been reported to produce ethylene, both in culture and in planta (Freebairn and Buddenhagen 1964; Weingart and Volksch 1997; Valls et al. 2006). These findings suggest that ethylene

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production by R. solanacearum and P. syringae plays an important role in pathogenesis. Consistent with this hypothesis, ethylene synthesis mutants of some strains of P. syringae pv. glycinea grow to significantly reduced levels in bean and soybean plants (Weingart et al. 2001). Interestingly, in addition to encoding the ethylene biosynthetic gene ACC oxidase, the R. solanacearum genome contains a gene encoding ACC deaminase (Salanoubat et al. 2002), an enzyme involved in ethylene degradation. This suggests that R. solanacearum may carefully modulate ethylene levels within the plant for a maximal virulence effect. The role of ethylene in interactions with bacterial plant pathogens is somewhat confusing, as it is reported to both suppress and stimulate host defenses responses, depending upon the nature of the interaction under study. For example, ethylene signaling is required for normal susceptible responses to P. syringae; ethylene-insensitive mutants (e.g., ein2 and ein3 eil1) are less susceptible to infection (Bent et al. 1992; Chen et al. 2009; Boutrot et al. 2010), and overexpression of ERF1, an ethylene-responsive transcription factor, results in increased susceptibility to P. syringae (Berrocal-Lobo et al. 2002). These observations are consistent with studies indicating that ethylene antagonizes SA signaling in these interactions (Chen et al. 2009). However, ethylene is also required for normal induction of PTI by the MAMP flg22. Challenge of Arabidopsis plants with flg22 results in increased ethylene production, and ethylene perception and signaling mutants are impaired for flg22-mediated responses (Boutrot et al. 2010). This effect appears to be mediated through the expression of the flg22 receptor, FLS2. Although ethylene plays multiple roles in plant-pathogen interactions, in the case of virulent pathogens such as P. syringae, it seems that the primary role of ethylene is to promote disease, as the basal defenses induced by MAMP perception are normally suppressed by pathogen virulence factors (van Loon et al. 2006).

Abscisic Acid Abscisic acid (ABA) is well known for the roles it plays in regulating seed development and germination and in mediating responses to abiotic stress induced by drought, salt, and cold (Hirayama and Shinozaki 2007). ABA and its downstream signaling components induce stomatal closure in response to drought and epiphytic bacteria, suggesting a common mechanism of guard cell response to both biotic and abiotic stresses (Melotto et al. 2006). However, the role of ABA in modulating post-invasion plant-bacterium interactions is not always clear. For instance, how ABA impacts the outcome of a pathogenic interaction appears to depend not only upon the lifestyle of the pathogen (i.e., whether it is a biotroph or a necrotroph) but also on the specific interaction (Fan et al. 2009; Robert-Seilaniantz et al. 2011). This may be due, in part, to the fact that ABA is involved in regulatory cross talk with the ethylene, JA, and SA defense signaling pathways (Bari and Jones 2009; Pieterse et al. 2009; Robert-Seilaniantz et al. 2011). More recent evidence suggests that ABA signaling may also impact pathogen growth by influencing physiological conditions in infected tissue.

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Several studies indicate that ABA promotes susceptibility to biotrophic pathogens, such as the bacterium P. syringae and the oomycete Hyaloperonospora arabidopsidis. Upon infection with virulent P. syringae, the ABA level increases in the host tissue (Fan et al. 2009; de Torres-Zabala et al. 2007). Additionally, overexpression of ABA biosynthetic genes (e.g., NCED5) and consequent overproduction of ABA render plants hypersusceptible to P. syringae and H. arabidopsidis, while mutant plants impaired in ABA biosynthesis or responsiveness exhibit reduced susceptibility (de Torres-Zabala et al. 2007; Fan et al. 2009). There is growing evidence that P. syringae modulates ABA signaling as a virulence strategy. The increase in ABA levels observed upon infection with P. syringae is correlated with the induction of ABA biosynthetic genes, and this appears to be dependent upon type three secretion effectors, such as AvrPtoB (de Torres-Zabala et al. 2007; Fan et al. 2009). Furthermore, at least one P. syringae effector, HopAM1, enhances both ABA sensitivity and pathogen virulence within the host. The effect of HopAM1 on pathogen virulence is most pronounced on drought-stressed plants, suggesting that one role for ABA in contributing to host susceptibility is by modulating water availability in infected tissue in a manner that favors pathogen growth (Goel et al. 2008; Beattie 2011). Fan et al. (2009) also demonstrated that the increase in ABA in infected tissue stimulated pathogen-induced JA accumulation. Thus, one mechanism by which ABA enhances pathogen growth appears to be through promoting the antagonistic cross talk between JA and SA signaling pathways, inhibiting defenses against biotrophic pathogens. However, the overall role of ABA in impacting the outcome of plant-pathogen interactions is clearly not that simple, as in some situations, ABA and JA appear to act antagonistically in modulating defense responses (de Torres-Zabala et al. 2009). Although some necrotrophic fungi, such as Botrytis cinerea and Fusarium oxysporum, produce ABA, we are not aware of any reports that bacterial pathogens can synthesize ABA. However, it would not be necessary for them to do so in order to modulate ABA signaling in the host, as this can be accomplished by stimulating ABA synthesis and/or by promoting relocalization within the plant or release from internal stores (RobertSeilaniantz et al. 2011).

Auxin The role of auxin in promoting plant cell division and growth in diseases caused by tumorigenic plant pathogens such as A. tumefaciens and P. savastanoi is well established (Pitzschke and Hirt 2010; Rodrı´guez-Palenzuela et al. 2010). Although, a number of non-gall-forming plant pathogens, including R. solanacearum, X. oryzae pv. oryzae, and several P. syringae strains, have been reported to produce indole acetic acid (IAA), the predominant naturally occurring active form of auxin, when grown in culture (Phelps and Sequeira 1968; Fett et al. 1987; Glickmann et al. 1998; Ansari and Sridhar 2000; Valls et al. 2006; Spaepen and Vanderleyden 2011), the involvement of this plant

growth regulator in disease caused by nontumorigenic bacterial pathogens is less well understood. Several P. syringae strains harbor an iaaL gene that encodes an enzyme believed to catalyze the conversion of IAA to IAA-lysine, a conjugated form of IAA that is believed to be biologically less active than free IAA (Glickmann et al. 1998). Thus, presumably these bacteria are not only able to produce auxin but are also able to adjust free IAA levels within the plant. IAA levels increase in Arabidopsis plants infected with virulent X. campestris or P. syringae strains (O’Donnell et al. 2003; Chen et al. 2007; Spaepen and Vanderleyden 2011). Although the source (i.e., host or pathogen) of this increase in free IAA has not been established, the fact that several Arabidopsis genes encoding enzymes involved either in IAA biosynthesis or in hydrolysis of IAA-amino acid conjugates are upregulated upon infection with P. syringae (Niyogi et al. 1993; Bartel and Fink 1994; Zhao and Last 1996; Hull et al. 2000; Tao et al. 2003; Thilmony et al. 2006; Kazan and Manners 2009) suggests that the increase in free IAA in infected plants is produced, at least in part, by the plant. There is growing evidence that this increase in free IAA levels is of benefit to the pathogen. For example, exogenous application of the auxin analogs 1-naphthaleneacetic acid (NAA) or 2,4-D to Arabidopsis plants resulted in increased disease susceptibility to P. syringae, while plant mutants or transgenic lines with impaired auxin signaling exhibit reduced susceptibility to P. syringae (Chen et al. 2007; Wang et al. 2007; Navarro et al. 2006). Thus, it is reasonable to propose that some plant pathogens may modulate free auxin levels within the plant as a strategy to promote pathogen growth and disease development. Three virulence factors that appear to contribute to this process have been identified: the P. syringae phytotoxin coronatine and two TTSS effector proteins, AvrRpt2 from P. syringae and AvrBs3 from X. campestris. For instance, the expression of several genes involved in either producing IAA or releasing free IAA from conjugated pools within the plant (e.g., IAR3) is induced by infection with wild-type DC3000 but not with coronatinedefective mutants (Thilmony et al. 2006; Uppalapati et al. 2005). Transgenic Arabidopsis plants constitutively expressing the P. syringae type III effector AvrRpt2 (and lacking the corresponding resistance gene, RPS2) exhibit enhanced sensitivity to auxin and accumulate elevated levels of free IAA (Chen et al. 2007), suggesting that AvrRpt2 may promote pathogen virulence by modulating host auxin physiology. Finally, AvrBs3 from X. campestris stimulates host cell enlargement, a process associated with auxin, and induces expression of a group of auxin-regulated genes (Marois et al. 2002). In susceptible pepper plants, delivery of AvrBs3 specifically induces the expression of a group of auxin-induced SAUR genes (Marois et al. 2002). However, unlike AvrRpt2, the presence of AvrBs3 does not appear to affect free IAA levels in infected plants (Marois et al. 2002). Thus, AvrBs3 may alter host auxin physiology by altering IAA responses downstream of free IAA production and release from internal pools. Several studies indicate that auxin promotes susceptibility by suppressing host defenses. For example, auxin downregulates

Virulence Strategies of Plant Pathogenic Bacteria

expression of defense-related genes in cultured tobacco cells and plant tissues (Shinshi et al. 1987; Rezzonico et al. 1998), and injection of auxin-producing A. tumefaciens or P. savastanoi bacteria into tobacco leaves prior to injection of an avirulent P. syringae strain inhibits the development of visible tissue collapse (i.e., the hypersensitive response or HR) associated with the host defense response. The ability of these strains to suppress the P. syringae-induced HR was dependent on the presence of functional auxin biosynthetic genes, suggesting that auxin is directly involved in suppressing the HR (Robinette and Matthysse 1990). Exogenous application of auxin represses induction of pathogenesis-related (PR) genes upon treatment with SA (Wang et al. 2007). Although not directly demonstrated, it is possible that IAA promotes susceptibility by suppressing SA-mediated defenses. Additional, although somewhat indirect, evidence that auxin promotes pathogenesis comes from studies of basal defenses responses. Navarro et al. (2006) demonstrated that flg22 treatment of Arabidopsis induces a microRNA (miR393) that targets several TIR1 family auxin receptor genes and thus inhibits auxin signaling. Likewise, Wang et al. (2007) demonstrated that SA inhibits auxin signaling and that this appears to be mediated through the repression of the TIR1 family of auxin receptors. Thus, one component of MAMP-mediated basal defense appears to involve suppression of auxin responses. Auxin may also promote pathogen virulence via SAindependent mechanism(s). For example, auxin may alter the physiology of host cells at the site of infection in a manner that could render host tissue more suitable for pathogen growth. Alternatively, or additionally, IAA or IAA-amino acid conjugates could impact the pathogen directly, for example, by regulating virulence gene expression (Yang et al. 2007). A recent study by Gonza´lez-Lamothea et al. (2012) shows that Botrytis cinerea and P. syringae infection promote the accumulation of an IAAamino acid conjugate, IAA-Asp, and that this compound directly impacts the pathogen by stimulating transcription of virulence genes.

The Complexities of Hormone Signaling Networks in Plant-Microbe Interactions Note that the amount of crosstalk between the hormone signaling pathways discussed above is significant (Gazzarrini and McCourt 2003; Pieterse et al. 2009; Grant and Jones 2009). For instance, after reading this chapter, it should be clear that a large amount of interplay exists between the auxin, ABA, SA, and JA signaling pathways, and pathogens such as P. syringae may take advantage of the mutually antagonistic crosstalk between these pathways to manipulate signaling within the plant (Kazan and Manners 2009, 2012; Spoel and Dong 2009; Grant and Jones 2009; Robert-Seilaniantz et al. 2011). Likewise, auxin appears to upregulate the expression of ACC synthase 4 (ACS4; Abel et al. 1995), an enzyme that catalyzes a rate-limiting step in ethylene biosynthesis. Thus, auxin may also induce ethylene biosynthesis. Moreover, increasing evidence suggests that auxin and JA

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signaling pathways are interconnected. Therefore, the involvement of one plant hormone in a plant-pathogen interaction could be mediated, at least in part, through the action of one or more other plant hormones. Future studies aimed at untangling these complicated signaling networks will undoubtedly provide valuable insight into the virulence mechanisms used by bacterial plant pathogens.

Challenges The recent use of a combination of genetic, molecular, and genomic approaches has led to major advances in the identification of numerous potential new virulence factors. Studies involving plant genetic, genomic, and biochemical approaches, as well as physiological and gene expression analyses of transgenic plants expressing specific pathogen virulence factors (e.g., type III effector proteins), have provided insights on the action of virulence factors in the host cell. However, despite many recent advances, little is known about the plant processes modulated by pathogens. The challenge that lies ahead is to continue to develop experimental strategies that will facilitate the investigation of the mode of action of these factors and how they function collectively within the plant to promote pathogen virulence and disease. Given the potential functional redundancy of these factors, and the fact that their mode of action may not always be accurately predicted, it would be wise to utilize a variety of approaches in these future studies. In certain situations, advantage can also be taken of the observations that certain bacterial virulence factors are active in yeast (for examples, see Abramovitch et al. (2003) and Jamir et al. (2004)). The power of yeast genetics and the use of heterologous systems will likely facilitate the identification of host components that are important in mediating the activity of these virulence factors. It is unclear how plants prioritize their response to multiple stresses they face in nature. Additional studies addressing the uniqueness and commonalities of regulatory networks controlling responses to abiotic and biotic stressors will shed some light on response output in the field and provide new tools for crop improvement. Collectively, these studies are likely to provide valuable information regarding the molecular mechanisms underlying pathogen virulence, the host processes that are modulated during pathogenesis, as well as how environmental inputs impact host-pathogen interactions. The insights gained from these experiments may also lead to the development of new approaches for controlling virulence and disease development in agronomically important plant-pathogen interactions.

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3 Oxidation of Inorganic Nitrogen Compounds as an Energy Source Eberhard Bock1 . Michael Wagner2 1 Institute of General Botany, Department of Microbiology, University of Hamburg, Hamburg, Germany 2 Department of Microbial Ecology, Faculty Center of Ecology, Faculty of Life Sciences, University of Vienna, Vienna, Austria

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Phylogeny of Lithotrophic Nitrifying Bacteria . . . . . . . . . . . . . 89 Phylogeny of Ammonia Oxidizers . . . . . . . . . . . . . . . . . . . . . . . 90 Phylogeny of Nitrite Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . 90 Biochemistry of Ammonia-Oxidizing Bacteria . . . . . . . . . . . . . 91 Ammonia and Hydroxylamine as Substrates . . . . . . . . . . . . . 93 Enzymes Involved in Ammonia Oxidation . . . . . . . . . . . . . . 93 Ammonia Monooxygenase . . . . . . . . . . . . . . . . . . . . . . . . . . . 93 Hydroxylamine Oxidoreductase . . . . . . . . . . . . . . . . . . . . . . 94 Electron Flow and Energy Transduction . . . . . . . . . . . . . . . . . 95 Electron Flow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Energy Transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 NADH Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 Co-oxidation and Inhibition of AMO . . . . . . . . . . . . . . . . . . . 96 Denitrification Catalyzed by Ammonia Oxidizers . . . . . . . 97 Genetics of Ammonia Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 Genes Encoding AMO, HAO, and Related Enzymes . . . . . 98 Regulation of AMO and HAO . . . . . . . . . . . . . . . . . . . . . . . . . . . 98 Biochemistry of Nitrite-Oxidizing Bacteria . . . . . . . . . . . . . . . . 99 Nitrite as a Substrate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 Enzymes Involved in Nitrite Oxidation . . . . . . . . . . . . . . . . . 100 Nitrite Oxidoreductase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 Cytochrome c Oxidase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 Nitrite Reductase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 101 Electron Flow and Energy Transduction . . . . . . . . . . . . . . . . 101 Electron Flow of the Conventional Respiratory Chain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 ATP Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 103 NADH Production and Cell Growth . . . . . . . . . . . . . . . . 103 Genetics of Nitrite Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 104 Heterotrophic Nitrification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Novel Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Nitrogenous Oxides Are Essential for Aerobic Ammonia Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 Anaerobic Ammonia Oxidation Catalyzed by Nitrosomonas eutropha . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

Anaerobic Ammonia Oxidation with Nitrogen Dioxide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108 Anaerobic Ammonia Oxidation in Cell-Free Extracts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108 Anaerobic Ammonium Oxidation Catalyzed by Deep Branching Planctomycetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Abstract This chapter covers one of the microbiological steps of the nitrogen cycle, nitrification, which is the biological oxidation of reduced forms of inorganic nitrogen to nitrite and nitrate. Nitrifying bacteria use the oxidation of inorganic nitrogen compounds as their major energy source. Reactions are catalyzed by two physiological groups of bacteria: ammonia-oxidizing bacteria, which gain energy from oxidation of ammonia to nitrite, and nitrite-oxidizing bacteria, which thrive by oxidizing nitrite to nitrate. Because of the toxic nature of nitrite, its rapid conversion to nitrate, assimilated by plants and microorganisms, is essential. Ammonia oxidizers are lithoautotrophic organisms using carbon dioxide as the main carbon source; ammonia monooxygenase oxidizes ammonia to hydroxylamine, which is converted to nitrite by the hydroxylamine oxidoreductase. When grown lithotrophically with nitrite, nitrite is oxidized to nitrate by the nitrite oxidoreductase and the oxygen atom in the nitrate molecule is derived from water. The enzyme also reduces nitrite to nitrate when Nitrobacter strains are grown heterotrophically in the presence of nitrate. Detailed schemes for electron flow and energy transduction as well as energy generation schemes are outlined and the role of nitrifying bacteria in the environment highlighted. The two groups of nitrifying bacteria are phylogenetically unrelated, as they are found in different classes of Proteobacteria and members of the nitrite oxidizers are even found in different phyla. This chapter also covers the physiology and phylogeny of recently detected anaerobic ammonium-oxidizing deep-branching members of the phylum Planctomycetes and of Nitrosomonas eutropha.

Introduction Life depends on the element nitrogen. In nature, nitrogen exists mainly in the oxidation states -III (NH3), O (N2), +I (N2O),

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_64, # Springer-Verlag Berlin Heidelberg 2013

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nitrogen fixation Air–N2

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NO2leaching

. Fig. 3.1 Nitrogen cycle mediated by the biosphere

+II (NO), +III (NO2 ), +IV (NO2), and +V (NO3 ). Owing to nitrogen transformations by the activity of living organisms and to chemical instability, any form of oxidation state has only a transient existence. Dinitrogen (N2) is the most inert and frequent constituent of the atmosphere. Taking into account also abiotic transformations, three cycles of nitrogen can be distinguished: 1. The cycle of the atmosphere 2. The interaction between the atmosphere and the biosphere 3. The cycle of the biosphere The nitrogen cycle mediated by the biosphere (> Fig. 3.1) can also be characterized by mobilization and immobilization of nitrogen compounds. Most of the reactions are catalyzed exclusively by prokaryotes. By microbial nitrogen fixation, dinitrogen is reduced to ammonia and subsequently transferred to amino acids and assimilated into cell material. On the other hand, ammonia is released from organic nitrogen compounds by microbial activity called ‘‘ammonification’’ or ‘‘mineralization.’’ Ammonia (NH3)/ammonium (NH4+) is the most frequently found form of nitrogen in the biosphere and is transferred efficiently over long distances via volatilization. In contrast, nitrite is usually found in trace amounts in aerobic habitats and only accumulates at low oxygen partial pressure, for example, in soil with high water potential. Because of the toxicity of nitrite for living organisms, the maintenance of low nitrite concentration in aerobic habitats is essential. Under oxic conditions, ammonia and nitrite are not stable and

are converted to nitrate by nitrifying bacteria. Nitrification, the biological oxidation of reduced forms of inorganic nitrogen to nitrite and nitrate, is catalyzed by two physiological groups of bacteria. Ammonia-oxidizing bacteria, which use ammonia and not ammonium as substrate (Suzuki et al. 1974), gain energy from oxidation of ammonia to nitrite, and nitrite-oxidizing bacteria thrive by oxidizing nitrite to nitrate. In seawater and freshwater as well as in soil, nitrite produced by the ammonia oxidizers is immediately consumed by nitrite oxidizers, and thus, the nitrite concentration is extremely low in these environments (El-Demerdash and Ottow 1983; Schmidt 1982). Nitrate can be assimilated by plants and microorganisms. Under anoxic or oxygen-limited conditions, nitrate is used as electron acceptor for anaerobic respiration (if organic matter is available) and thereby converted to ammonia (respiratory ammonification) or dinitrogen (denitrification). This chapter focuses on nitrifying bacteria, which use the oxidation of inorganic nitrogen compounds as their major energy source. Lithotrophic nitrifiers are Gram-negative bacteria and conventionally have been placed in the family Nitrobacter iaceae (Buchanan 1917; Watson 1971; Watson et al. 1989). However, phylogenetically the lithoautotrophic ammonia oxidizers, characterized by the prefix Nitroso-, and nitrite oxidizers, characterized by the prefix Nitro-, are not closely related (Teske et al. 1996; Purkhold et al. 2000). Comparative 16S rRNA sequence analysis demonstrated that all recognized ammonia oxidizers are either members of the b- or g-subclass of Proteobacteria (> Fig. 3.2). The genera

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

Nitrosospira briensis Nitrosovibrio tenuis Nitrosospira multiformis Nitrosomonas cryotolerans Nitrosomonas ureae Nitrosomonas oligotropha Nitrosomonas nitrosa Nitrosomonas communis Nitrosomonas aestuarii Nitrosomonas marina Nitrosomonas halophila Nitrosomonas europaea Nitrosomonas eutropha Nitrosococcus mobilis Zoogloea ramigera

0.10

Arhodomonas aquaeolei Nitrococcus mobilis Ectothiorhodospira halophila Ectothiorhodospira shaposhnikovi Nitrosococcus halophilus Nitrosococcus oceani Escherichia coli Nitrobacter alkaticus Nitrobacter winogradskyi Nitrobacter vulgaris “Nitrobacter hamburgensis” Rhodopseudomonas palustris Bradyrhizobium japonicum Afipia clevelandensis Nitrospina gracilis Bdellovibrio bacteriovorus Pelobacter propionicus Desulfobacter postgatei Nitrospira marina Nitrospira moscoviensis Leptospirillum ferrooxidans Thermodesulfovibrio islandicus “Magnetobacterium bavaricum”

3

Beta Proteobacteria

Gamma Proteobacteria

Alpha Proteobacteria

Delta Proteobacteria

Nitrospira-Phylum

. Fig. 3.2 16S rRNA-based tree reflecting the phylogenetic relationship of ammonia- and nitrite-oxidizing bacteria. The consensus tree is based on the results of a maximum likelihood analysis of the 16S rRNA primary structure data from the nitrifying bacteria shown in the tree and a selection of reference sequences. Only homologous positions that share identical residues in at least 50 % of all available almost complete bacterial 16S rRNA sequences were included for tree reconstruction. In the tree, ammonia oxidizers are labeled green, and nitrite oxidizers are depicted in red. It should be noted that the assignment of the genus Nitrospina to the d-Proteobacteria is tentative and might change if additional reference sequences become available. Multifurcations connect lineages for which no unambiguous branching order could be retrieved using different treeing methods. Bar represents 10 % estimated sequence divergence

Nitrosomonas (including Nitrosococcus mobilis), Nitrosospira, Nitrosolobus, and Nitrosovibrio form a closely related monophyletic assemblage within the b-subclass of Proteobacteria (Head et al. 1993; Woese et al. 1984; Teske et al. 1994; Uta˚ker et al. 1995; Pommerening-Ro¨ser et al. 1996; Purkhold et al. 2000), whereas the genus Nitrosococcus constitutes a separate branch within the g-subclass of Proteobacteria (Woese et al. 1985; Purkhold et al. 2000). Among the nitrite oxidizers, the genera Nitrobacter, Nitrococcus, and Nitrospina

were assigned to the a-, g-, and g-subclass of Proteobacteria, respectively (Orso et al. 1994; Teske et al. 1994). Nitrite oxidizers of the genus Nitrospira are affiliated with the recently described Nitrospira phylum, which represents an independent line of descent within the domain Bacteria (Ehrich et al. 1995). The most important character of lithotrophic nitrifying bacteria is energy generation via ammonia oxidation to nitrite (section > ‘‘Biochemistry of Ammonia-Oxidizing Bacteria’’) and nitrite oxidation to nitrate (section > ‘‘Biochemistry of

85

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Oxidation of Inorganic Nitrogen Compounds as an Energy Source

Nitrite-Oxidizing Bacteria’’), respectively, according to the following equations: NH3 þ 0:5O2 ! NH2 OH

ð3:1Þ

1

DG0 0 ¼ þ17 kJ
mol

0:5O2 þ 2Hþ þ 2e ! H2 O DG0 0 ¼

137 kJ
mol

ð3:2Þ

1

NH3 þ O2 þ 2Hþ þ 2e ! NH2 OH þ H2 O DG0 0 ¼

120 kJ
mol

1

ð3:3Þ

1

ð3:4Þ

NH2 OH þ H2 O ! HNO2 þ 4Hþ þ 4e DG0 0 ¼ þ23 kJ
mol 0:5O2 þ 2Hþ þ 2e ! H2 O DG0 0 ¼

137 kJ
mol

ð3:5Þ

1

NH2 OH þ 1:5O2 ! HNO2 þ 2e þ 2Hþ DG0 0 ¼

114 kJ
mol

NH3 þ 1:5O2 ! HNO2 þ H2 O DG0 0 ¼

235 kJ
mol

NO2 þ H2 O ! NO3 þ 2Hþ þ 2e

0:5O2 þ 2Hþ þ 2e ! H2 O DG0 0 ¼

137 kJ
mol

NO2 þ 0:5O2 ! NO3 DG0 0 ¼ > Equations

54 kJ
mol

1

1

ð3:6Þ

ð3:7Þ

1

DG0 0 ¼ þ83 kJ
mol

1

1

ð3:8Þ

ð3:9Þ

ð3:10Þ

3.1 and > 3.2 describe the two half-reactions of ammonia oxidation to the intermediate hydroxylamine (NH2OH). The total reaction is given in > Eq. 3.3. For hydroxylamine oxidation, no oxygen is consumed (> Eq. 3.4). Subsequently two electrons are transferred back to reaction > 3.2, and the remaining two electrons pass to the respiratory chain (> Eq. 3.5). The second step of ammonia oxidation, the hydroxylamine oxidation, is depicted in > Eq. 3.6. The overall reaction (> Eq. 3.7) shows that biogenic ammonia oxidation causes nitric acid production. The dG00 value of reaction > 3.7 is significantly higher than that of nitrite oxidation (> Eq. 3.10). Nitrite oxidation starts with > Eq. 3.8. Electrons are released and penetrate the respiratory chain at the cytochrome c level (> Eq. 3.9). There is no acid production when nitrite is oxidized to nitrate (> Eq. 3.10). Ammonia oxidation is initiated by the enzyme ammonia monooxygenase (AMO; sections > ‘‘Enzymes Involved in Ammonia Oxidation’’; > ‘‘Ammonia Monooxygenase’’), which oxidizes ammonia to hydroxylamine. Substrates for AMO are

ammonia (Wood 1986), dioxygen, and two electrons. One atom of molecular oxygen is reduced to water, while the second oxygen atom is incorporated to form hydroxylamine. The intermediate hydroxylamine is further oxidized to nitrite by hydroxylamine oxidoreductase (HAO; sections > ‘‘Enzymes Involved in Ammonia Oxidation’’; > ‘‘Hydroxylamine Oxidoreductase’’). Two of the four electrons derived are required for AMO activity, and the other two are used for energy generation (section > ‘‘Electron Flow and Energy Transduction’’). The AmoA protein is assumed to contain the active site of AMO (Hyman and Arp 1992). A second AMO subunit named ‘‘AmoB’’ has been identified (Bergmann and Hooper 1994a). The gene cluster encoding AMO contains a third open reading frame termed ‘‘amoC,’’ which is located upstream of the genes amoA and amoB (Klotz et al. 1997; section > ‘‘Genetics of Ammonia Oxidizers’’). Neither AmoA nor AmoB has been purified in the active state as yet. The enzyme HAO is a trimer of 63-kDa subunits, including seven c-type hemes and a novel heme (P-460) per monomer (Arciero and Hooper 1993; Hoppert et al. 1995; Igarashi et al. 1997). The enzyme is located in the periplasmic space but anchored in the cytoplasmic membrane. Nitrite oxidation is initiated by the enzyme nitrite oxidoreductase (NO2-OR; section > ‘‘Enzymes Involved in Nitrite Oxidation’’) which occurs as characteristic membrane-associated two-dimensional crystals in all nitrite oxidizers. These regularly arranged particles are located on the surface of the cytoplasmic—and if present—intracytoplasmic membranes of nitriteoxidizing bacteria. In all Nitrobacter species and in Nitrococcus, particles are arranged in rows, whereas in Nitrospina and both Nitrospira species, hexagonal patterns were observed. The NO2-OR consists of two subunits (Meincke et al. 1992). For Nitrobacter hamburgensis, the molecular weight of one particle was found to be 186 kDa representing an ab-heterodimer (Spieck et al. 1996). The full sequence of the b-subunit as well as a partial sequence of the a-subunit of the NO2-OR of Nitrobacter hamburgensis shows similarities to nitrate reductases of several chemoorganotrophic bacteria (section > ‘‘Genetics of Nitrite Oxidizers’’). During oxidation of nitrite to nitrate, the additional oxygen atom of nitrate is derived from water (Aleem 1965) and two electrons are released for energy generation. In addition to lithotrophic nitrifiers, various heterotrophic bacteria, fungi, and algae (Focht and Verstraete 1977; Killham 1986; Papen et al. 1989) are capable of oxidizing ammonia to nitrate. However, in contrast to lithotrophic nitrification, heterotrophic nitrification is not coupled to energy generation (section > ‘‘Heterotrophic Nitrification’’). Consequently, heterotrophic nitrifiers are dependent on the oxidation of organic substrates (Focht and Verstraete 1977; Kuenen and Robertson 1987). During heterotrophic nitrification, ammonia or reduced nitrogen from organic compounds (e.g., the amino group of amino acids) is co-oxidized to hydroxylamine, gaseous nitrogen oxides, nitrite, or nitrate. For example, methane-oxidizing bacteria were shown to co-oxidize ammonia to nitrite by a biochemically well-characterized particulate (membranebound) methane monooxygenase and a unique hydroxylaminoxidoreductase (Anthony 1982; Yoshinari 1985; O’Neil and

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

Wilkinson 1977; Zahn et al. 1994; Bergmann et al. 2000). The methane monooxygenase is assumed to be biochemically related to the AMO of ammonia-oxidizing bacteria, and methaneoxidizing bacteria are potential contributors to nitrification in the rhizosphere of rice plants (Bodelier and Frenzel 1999). Conversely, ammonia oxidizers are able to oxidize methane to methanol (Hyman and Wood 1983; Ward 1987; Jones and Morita 1983; Steudler et al. 1996), but up to now, there is no evidence that ammonia oxidizers significantly contribute to the oxidation of atmospheric methane (CH4) in natural systems (Jlang and Bakken 1999; Bodelier and Frenzel 1999). In general, heterotrophic nitrification is considered to contribute only marginally to the global nitrogen cycle (Brady 1984; Brown 1988) but nevertheless might be of local importance especially in heath and conifer forest soils (e.g., see Van de Dijk and Troelstra 1980; Schimel et al. 1984). Lithotrophic nitrifiers are autotrophic bacteria that fix carbon dioxide (CO2) via the Calvin-Benson cycle (Harms et al. 1981) and, to a lesser extent, via phosphoenolpyruvate carboxylase (Takahashi et al. 1993). In the past, they were thus described as obligate lithoautotrophs and were thought to find organic compounds toxic. However, this assumption is not correct for several nitrifier species. Clark and Schmidt (1967) demonstrated that ammonia oxidizers of the genus Nitrosomonas and nitrite oxidizers of the genus Nitrobacter are capable of growing mixotrophically with ammonia or nitrite as electron donors and with a combination of carbon dioxide and organic compounds as carbon source. Compared to purely autotrophic growth, the addition of organic compounds stimulated cell growth and increased cell yield (Steinmu¨ller and Bock 1976; Matin 1978; Kru¨mmel and Harms 1982; Watson et al. 1986). Furthermore, the nitrite oxidizers Nitrobacter winogradskyi, N. hamburgensis, and N. vulgaris can grow chemoorganotrophically with acetate or pyruvate as electron donor and dioxygen or nitrate (in absence of dioxygen) as electron acceptor (Bock 1976; Freitag et al. 1987). However, for these organisms, heterotrophic growth was always slower than lithotrophic growth. Recently, Daims and coworkers (Daims et al. 2000, 2001) showed that in nitrifying activated sludge, not yet cultured Nitrospira-related nitrite oxidizers fix CO2 and simultaneously take up pyruvate but not acetate, butyrate, and propionate. In addition, some strains of Nitrosomonas can utilize organic substances like urea or glutamine as source of their substrate ammonia for lithotrophic growth (Koops et al. 1991). The transformation of ammonia to nitrate via nitrite by the nitrifying bacteria has various direct and indirect implications for natural and man-made systems. For example, nitrifying bacteria contribute directly or indirectly to loss of nitrogen compounds from various environments due to: 1. Leaching of mobile nitrogen compounds produced by nitrifiers. Leaching is the mobilization and transfer of nitrate to rivers, lakes, seawater, and groundwater. Nitrification is not desirable in agricultural soil because it induces loss of soil nitrogen. Fertilizer ammonium, which is required for

3

plant growth, adsorbs well to clay particles of soil owing to its positive charge (> Fig. 3.1). When converted to nitrate, the inorganic soil nitrogen becomes mobile and thus susceptible to denitrification and leaching. In some countries, nitrification inhibitors like nitrapyrin (N-Serve) are used in agriculture to minimize nitrogen loss (Huber et al. 1977; Keeny 1986; Slangen and Kerkhoff 1984; Lipschultz et al. 1981; Poth and Focht 1985). 2. Denitrification. Denitrification is the microbial reduction of nitrate via nitrite, nitric oxide (NO) and nitrous oxide (N2O) to dinitrogen (N2). This type of anaerobic respiration can be performed by a variety of phylogenetically different heterotrophic microorganisms. By aerobic oxidation of ammonia to nitrate, nitrifying bacteria produce the electron acceptor for subsequent denitrification in many natural and engineered systems. During the last years, nitrifiers also have been shown to be able to denitrify (section > ‘‘Denitrification Catalyzed by Ammonia Oxidizers’’). 3. Chemodenitrification of nitrite (produced by the ammonia oxidizers) in acidic environments. Chemodenitrification is defined as nonenzymatically catalyzed loss of nitrogen due to dismutation of nitric acid at pH values ‘‘Anaerobic Ammonium Oxidation Catalyzed by Deep Branching Planctomycetes’’ in this chapter). Nitrifying bacteria are slow-growing organisms because their cell growth is inefficient. For example, nitrite oxidizers oxidize 85–115 mol of nitrate to generate the energy required for assimilation of 1 mol of carbon dioxide (Bo¨meke 1954). Thus, it is not surprising that the shortest generation times measured in laboratory experiments did not exceed 7 h for Nitrosomonas and 10 h for Nitrobacter (Bock et al. 1990). For cell division in natural environments, most nitrifier species even need several days to weeks depending on substrate, oxygen availability, the temperature, and pH values. The slow growth rates of nitrifiers have severely hampered cultivation-dependent approaches to investigate the number, community composition, and dynamics of nitrifiers in different environments. The number of nitrifiers in complex systems has been traditionally determined by the most probable number (MPN) technique (Matulewich et al. 1975). However, this method is timeconsuming, and the nitrifier cell counts determined usually do not correlate well with nitrifying potential estimated for the same environmental sample under optimized laboratory conditions (Belser and Mays 1982; Belser 1979; Groffmann 1987; Mansch and Bock 1998). These discrepancies illustrate that not all nitrifiers can be cultivated using standard methods (Stephen et al. 1998; Juretschko et al. 1998; Purkhold et al. 2000). Furthermore, in many environments, nitrifiers form dense microcolonies of ten to several thousand cells embedded in extracellular polymeric substances (EPS; > Fig. 3.3). Since these microcolonies are resistant to the dispersal techniques implemented in standard cultivation protocols, the use of these protocols dramatically underestimates the number of nitrifiers occurring in microcolonies (Watson et al. 1989; Stehr et al. 1995; Wagner et al. 1995).

a

b

10.000x

30.000x

. Fig. 3.3 Transmission electron micrographs of ultrathin sections of an ammonia oxidizer microcolony in activated sludge (a) Arrows indicate intracytoplasmic membranes (b) Modified from Wagner et al. (1995)

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

For direct microscopic enumeration of nitrifiers in complex samples, the fluorescent antibody (FA) technique can be applied (Belser 1979; Fliermanns et al. 1974), but for antibody production, the target cells have to be isolated first as pure culture and the produced antibodies often recognize only a few strains of a species (Belser and Schmidt 1978). Recently monoclonal antibodies targeting the nitrite oxidoreductase were developed that allow group-specific detection of nitrite-oxidizing bacteria (Bartosch et al. 1999). In addition, polyclonal antibodies specifically recognizing the AmoB protein of b-subclass ammonia oxidizers are available (Pinck et al. 2001). Alternatively, nitrifiers can be detected in environmental samples independent from their culturability by using a variety of different polymerase chain reaction (PCR) techniques for specific amplification of 16S rRNA gene fragments (e.g., Degrange and Bardin 1995; Hiorns et al. 1995; Voytek and Ward 1995; McCaig et al. 1994; Kowalchuk et al. 1997; Uta˚ker and Nes 1998) or a fragment of the amoA gene (e.g., Rotthauwe et al. 1997; Purkhold et al. 2000). Quantitative population structure analysis of nitrifying bacteria within their natural habitat can most precisely be obtained by applying the recently developed set of rRNAtargeted oligonucleotide probes for fluorescence in situ hybridization (FISH; Wagner et al. 1995, 1996; Mobarry et al. 1996; Juretschko et al. 1998; Daims et al. 2000; > Fig. 3.4). Nitrifying bacteria are present in oxic and even anoxic environments. They are widely distributed in freshwater, seawater,

3

soils, on/in rocks, in masonry, and in wastewater treatment systems. Nitrifiers also could be enriched or isolated from extreme habitats like heating systems with temperatures of up to 47  C (Ehrich et al. 1995; E. Lebedeva, personal communication) and permafrost soils up to a depth of 60 m at a temperature of down to 12  C. Although the pH optimum for cell growth is 7.6–7.8, nitrifiers were frequently detected in environments with suboptimal pH (e.g., acid tea soils and forest soils at pH values below 4) but also in highly alkaliphilic soda lakes at a pH of 9.7–10.5 (Sorokin et al. 2001). Growth under suboptimal acidic conditions might be possible by ureolytic activity, by aggregate formation (De Boer et al. 1991), or as biofilms (e.g., on clay particles; Allison and Prosser 1993). In many environments, nitrifier sensitivity to sunlight is of ecological importance. The light sensitivity of ammonia and nitrite oxidizers increases from blue light to long wave UV (Hooper and Terry 1974; Hyman and Wood 1984a; Shears and Wood 1985). Based on spectroscopic similarities, Shears and Wood (1985) postulated a model of the ammonia monooxygenase light inhibition similar to the three-stage catalytic cycle of the tyrosinase reaction. In Nitrobacter, which is more sensitive to visible light than Nitrosomonas (Bock 1965), the photooxidation of c-type cytochromes is assumed to cause light-induced cell death (Bock 1970). Although Nitrosomonas europaea and Nitrobacter sp. are the most commonly investigated ammonia and nitrite oxidizers in laboratory studies, molecular analysis revealed that other nitrifiers are of higher importance in many natural and engineered systems. For example, stone material of historical buildings and many soil systems seem to be dominated by members of the genera Nitrosovibrio and Nitrosospira, respectively (Spieck et al. 1992; Hiorns et al. 1995; Stephen et al. 1996; Meincke et al. 1989), whereas different Nitrosomonas species and Nitrosococcus mobilis are the most abundant ammonia oxidizers in wastewater treatment plants (Juretschko et al. 1998; Purkhold et al. 2000). Interestingly, not yet cultured members of the genus Nitrospira and not Nitrobacter are the most abundant nitrite oxidizers in sewage treatment plants and aquaria filters (Burrell et al. 1998; Juretschko et al. 1998; Wagner et al. 1996; Daims et al. 2000).

Phylogeny of Lithotrophic Nitrifying Bacteria

. Fig. 3.4 In situ detection (with fluorescently labeled 16S rRNA-targeted oligonucleotide probes) of ammonia-oxidizing and nitrite-oxidizing bacteria in a nitrifying biofilm from a municipal wastewater treatment plant. Ammonia oxidizers are stained red, whereas nitrite-oxidizing bacteria of the genus Nitrospira appear green. Bar = 10 mm

Traditionally, nitrifying bacteria have been lumped together into one coherent group, the family Nitrobacter iaceae (Watson 1971; Watson et al. 1989). Based on their ability to lithotrophically oxidize either ammonia to nitrite or nitrite to nitrate, nitrifying bacteria were separated into two groups, the ammonia and the nitrite oxidizers. The assignment of ammonia- and nitriteoxidizing bacteria into genera was dependent primarily upon their morphological features like cell size, shape, and the arrangement of the intracytoplasmic membranes (Watson et al. 1989). The physiological and morphological grouping of the nitrifying bacteria is in contradiction to data obtained from molecular phylogenetic studies which show at least subdivision

89

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3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

level diversity within and between the ammonia and nitrite oxidizers (Head et al. 1993; Orso et al. 1994; Teske et al. 1994; Purkhold et al. 2000; Ehrich et al. 1995). Significant differences between ammonia- and nitrite-oxidizing bacteria are also indicated by the fact that both physiological groups possess very different key enzyme systems for the energy-gaining oxidation of ammonia and nitrite, respectively (sections > ‘‘Enzymes Involved in Ammonia Oxidation’’; > ‘‘Enzymes Involved in Nitrite Oxidation’’). With the exception of the nitrite oxidizers of the genera Nitrospina and Nitrospira, all known nitrifiers are closely related to phototrophs and thus presumably originated in several independent events by conversion of photosynthetic ancestors to chemolithotrophs (Teske et al. 1994). Consistent with this conversion hypothesis, all nitrifying bacteria related to phototrophs retain the general structural features of the putative ancestor’s photosynthetic membrane complex, while nitrite oxidizers of the genera Nitrospina and Nitrospira lack intracytoplasmic membranes (ICMs). However, it should be noted that the ammonia oxidizers of the genera Nitrosospira and Nitrosovibrio lack an extensive intracytoplasmic membrane system (Koops and Mo¨ller 1992).

Proteobacteria, most closely related to the iron oxidizer Gallionella ferruginea. This lineage encompasses the genera Nitrosomonas (including Nitrosococcus mobilis, which is actually a member of the genus Nitrosomonas), Nitrosovibrio, Nitrosolobus, and Nitrosospira. It has been suggested (Head et al. 1993) and subsequently questioned (Teske et al. 1994) that the latter three genera should be reclassified into the single genus Nitrosospira. The nitrosomonads can be further subdivided into the N. europaea/Nc. mobilis cluster, the N. marina cluster, the N. oligotropha cluster, and the N. communis cluster (Purkhold et al. 2000). Nitrosomonas cryotolerans forms a separate lineage within the b-Proteobacteria. The genera Nitrosospira, Nitrosolobus, and Nitrosovibrio are closely related and form a cluster to the exclusion of the nitrosomonads. Similar but not identical evolutionary relationships were obtained if comparative analysis of AmoA sequences were performed (Purkhold et al. 2000). In the AmoA tree, the N. europaea/Nc. mobilis cluster, the N. marina cluster, and the Nitrosospira cluster are retained, whereas the members of the N. oligotropha cluster and the N. communis cluster form no monophyletic assemblages.

Phylogeny of Ammonia Oxidizers

Phylogeny of Nitrite Oxidizers

Chemolithotrophic ammonia oxidizers were isolated for the first time at the end of the nineteenth century (Winogradsky 1892). Since then, 16 species of ammonia oxidizers have been described (Jones et al. 1988; Koops et al. 1976, 1990; 1991, Watson 1965), and according to DNA-DNA hybridization experiments, at least 15 additional genospecies are ‘‘hidden’’ in existing culture collections (Koops et al. 1991; Koops and Harms 1985; Stehr et al. 1995). Our current perception of evolutionary relationships of ammonia-oxidizing bacteria is mainly based on comparative sequence analysis of their genes encoding the 16S rRNA and the active site polypeptide of the ammonia monooxygenase (AmoA). During the last decade, the genes for both biopolymers were sequenced for all recognized ammonia oxidizer species (Alzerreca et al. 1999; Head et al. 1993; Pommerening-Ro¨ser et al. 1996; Teske et al. 1994; Purkhold et al. 2000; Rotthauwe et al. 1995, 1997; McTavish et al. 1993; Horz et al. 2000), and the deduced phylogeny now provides an encompassing and relatively robust framework for assignment of 16S rDNA and amoA sequences of (1) ammonia oxidizer isolates (Stehr et al. 1995; Suwa et al. 1997; Uta˚ker et al. 1995; Juretschko et al. 1998) and (2) cloned sequence fragments directly retrieved from the environment (e.g., Stephen et al. 1996; Rotthauwe et al. 1995; Purkhold et al. 2000). According to comparative 16S rRNA sequence analysis, all recognized ammonia oxidizers are members of two monophyletic lineages within the b- and g-subclass of Proteobacteria (> Fig. 3.5). The marine species Nitrosococcus halophilus and Nitrosococcus oceani, which are distantly related to methaneoxidizing bacteria, cluster together in the g-subclass of Proteobacteria. All other ammonia oxidizers form a monophyletic assemblage within the b-subclass of

Four different genera, Nitrobacter, Nitrococcus, Nitrospina, and Nitrospira, of lithotrophic nitrite-oxidizing bacteria have been described. From 16S rRNA sequence analysis, the first three genera were assigned to different subclasses of the Proteobacteria, whereas Nitrospira is the name-giving genus of an independent bacterial phylum (> Fig. 3.5). The genus Nitrobacter contains the four closely related species (N. hamburgensis, N. vulgaris, N. winogradskii, and N. alkalicus) within the a-subclass of Proteobacteria. Nitrite oxidizers of the genus Nitrobacter are phylogenetically related to Bradyrhizobium japonicum, Blastobacter denitrificans, Afipia felis, Afipia clevelandensis, and the phototroph Rhodobacter palustris (Seewaldt et al. 1982; Orso et al. 1994; Teske et al. 1994) with which Nitrobacter shares a nearly identical arrangement of ICMs. The genus Nitrococcus represented by the single marine species Nitrococcus mobilis is, like the marine ammonia oxidizers of the genus Nitrosococcus, a member of the ectothiorhodospira branch of the g-subclass of Proteobacteria, consistent with an assumed photosynthetic ancestry of these nitrifiers. Nitrococcus and Nitrosococcus are the only nitrite and ammonia oxidizers that are relatively closely related, but the closest relatives of Nitrococcus mobilis are the phototrophic bacteria Arhodomonas aquaeoli, Ectthiorhodospira halochloris, and Ectthiorhodospira halophila (Teske et al. 1994). The genus Nitrospina with the marine Nitrospina gracilis as the only species (represented by two isolates, one from the Atlantic and the other from the Pacific) has been provisionally assigned to the d-subclass of Proteobacteria and is the only member of a deep branch within this subclass (Teske et al. 1994). Nitrospina gracilis shows no ICMs.

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

3

16S rRNA Beta AOB N. europaea/Nc. mobilis cluster N. marina cluster N. eutropha Nitrosomonas Nm63 N. Nitrosomonas Nm51 aestuarii 97 N. marina 100

N. oligotropha cluster N. oligotropha

98 92 45

63 99

67 100

N. ureae N. cryotolerans

100 100

N. communis N. nitrosa cluster Nitrosomonas Nm41

N. halophila

96

N. europaea

100 N. europaea Nm103

Nm107 Nc. mobilis Nc2 Nm104 Nc. mobilis Nm93 98

100

100

100 97

N. communis

75 100

Nitrosospira cluster

Comamonas testosteroni

to outgroups

Nitrosococcus oceani C-107T, C-27 Nitrosococcus sp. C-113

Gamma AOB

Methylomonas methanica Methylomicrobium album

100

39

Nitrosomonas Nm33

Nitrosococcus halophilus Nc4

58

79

Methylobacter sp. BB5.1

100 80

Methylocaldum tepidum Methylocaldum gracile Methylocaldum szegediense Methylococcus capsulatus

Gamma MOB

Sphaerotilus natans

Beta non AOB

0.10

. Fig. 3.5 Phylogenetic neighbor-joining 16S rRNA tree reflecting the relationships of ammonia-oxidizing bacteria and several reference organisms. The multifurcation connects branches for which a relative order could not be unambiguously determined by applying different treeing methods. Parsimony bootstrap values for branches are reported. Missing bootstrap values indicate that the branch in question was not recovered in the majority of bootstrap replicates by the parsimony method. AOB ammonia-oxidizing bacteria, MOB methane-oxidizing bacteria. The bar indicates 10 % estimated sequence divergence (Modified from Purkhold et al. 2000)

The genus Nitrospira encompasses the marine species Nitrospira marina and Nitrospira moscoviensis, isolated from a municipal water heating system. The genus Nitrospira forms a monophyletic grouping with the genera Thermodesulfovibrio, Leptospirillum, and with ‘‘Magnetobacterium bavaricum.’’ This phylogenetic assemblage has recently been identified as a novel phylum within the domain Bacteria and was named ‘‘Nitrospira phylum’’ (Ehrich et al. 1995). There is accumulating molecular evidence that Nitrospira-related nitrite oxidizers are of major importance for nitrite oxidation in wastewater treatment plants and aquarium filters (Burrell et al. 1998; Juretschko et al. 1998; Hovanec et al. 1998; Daims et al. 2000) and also occur in many natural environments including the rhizosphere (> Fig. 3.6). Like Nitrospina gracilis, members of the genus Nitrospira do not possess ICMs and are apparently not closely related to phototrophic bacteria.

Biochemistry of Ammonia-Oxidizing Bacteria Ammonia oxidizers are lithoautotrophic organisms using carbon dioxide as the main carbon source (Bock et al. 1991). Their

only way to gain energy is the two-step oxidation of ammonia to nitrite (Hooper 1969). Investigations of the Km values and pH optima indicate that ammonia (NH3) rather than ammonium (NH4+) is the substrate of ammonia oxidizers (Suzuki et al. 1974; Drozd 1976). This is in accordance with results showing that the ammonia-oxidizing enzyme might be located in the cytoplasmic membrane (Suzuki and Kwok 1981; Tsang and Suzuki 1982), since membranes are highly permeable to ammonia but not to ammonium (Kleiner 1985). First, ammonia is oxidized to hydroxylamine (Kluyver and Donker 1926) by the ammonia monooxygenase (AMO; Hollocher et al. 1981). This enzyme does not possess high substrate specificity and also oxidizes several apolar compounds such as methane, carbon monoxide, or some aliphatic and aromatic hydrocarbons (Hooper et al. 1997). These compounds can act as competitive inhibitors of ammonia oxidation (Hyman et al. 1988; Keener and Arp 1993). The second step is performed by the hydroxylamine oxidoreductase (HAO). This enzyme oxidizes hydroxylamine to nitrite (Wood 1986). Two of the four electrons released (Andersson and Hooper 1983) are required for the AMO reaction (Tsang and Suzuki 1982), whereas the remaining ones are used for the generation of proton motive force

91

92

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

0.10

six SBBR clones activated sludge clone A1-11 (AF033558) activated sludge clone A1-4 (AF033559) SBR clone RC73 (v14641) SBR clone RC90 (Y14642) SBR clone RC11 (Y14636) SBR clone RC99 (Y14643) SBR clone RC7 (Y14640) SBR clone RC14 (Y14637) SBR clone SBR2016 (X84560) SBR clone RC25 (Y14639) SBR clone RC19 (Y14638) fluidized bed reactor clone b30 (AJ224043) fluidized bed reactor clone b2 (AJ224038) fluidized bed reactor clone b21 (AJ224043) fluidized bed reactor clone b9 (AJ224045) fluidized bed reactor clone g6 (AJ224039) fluidized bed reactor clone o14 (AJ224044) fluidized bed reactor clone o9 (AJ224042) SBR clone SBR2015 (X84460) SBR clone SBR2046 (X84584) SBR clone SBR1024 (X84468) activated sludge clone GC86 (Y14644) soil done (AF 010059) SBR clone SBR2046 (AF 155155) mesotrophic lake done LCo23 (AF337199) Lale Baikal done 1405-19 (AJ007652) rhizosphere done (AJ232865) rhizosphere done (AJ232865) rhizosphere done RSC-11-52 (AJ252684) freshwater done MNC 2 (AF 293010) Nitrospira moscoviensis (X82558) clone C26-12 (AF332345) aquarium clone (AF035813) aquarium clone WJGRT-122 (AF175639) SBR clone SBR1065 (X84499) soil clone (AF010096) Nullarbor caves clone wb1 CO2 (AF317758)

I

II

done T26-17 (AF 332306) done C26-14 (AF 332330) soil done C059 (AF 128729) Nullarbor caves clone wb1_N07 (AF317779) Nullarbor caves clone wb1_K02 (AF317775) Nullarbor caves clone wb1_K22 (AF317777) Nullarbor caves clone wb1_K07 (AF317764) done T26-16 AF 332296) Nullarbor caves clone wb1_C17 (AF317762)

III

deltaicrnud done n4r (AF 194185) Nitrospira marina (X82559) deep sea clone BD 3-8 (AB015550)

IV

sponge symbiont WS65 (AF 18451) sponge symbiont WS69 (AF 186454) sponge symbiont WS58 (AF 186446) Nullarbor caves clone wb1_B21 (AF317752)

. Fig. 3.6 Phylogenetic tree of the genus Nitrospira based on comparative analysis of 16S rRNA sequences. The basic tree topology was determined by maximum likelihood analysis of all sequences longer than 1,300 nucleotides. Shorter sequences were successively added without changing the overall tree topology. Branches leading to sequences shorter than 1,315 nucleotides are dotted to point out that the exact affiliation of these sequences cannot be determined. Black spots on tree nodes symbolize high parsimony bootstrap support above 90 % based on 100 iterations. The scale bar indicates 0.1 estimated changes per nucleotide. The four sublineages of the genus Nitrospira are delimited by horizontal dashed lines and marked by the numbers I to IV. Two of the four sublineages entirely consist of 16S rDNA sequences amplified from environmental samples (Modified from H. Daims et al. 2001)

(Hollocher et al. 1982) to regenerate ATP and NADH (Wheelis 1984; Wood 1986). Most of the investigations on energy metabolism of ammonia-oxidizing bacteria have been carried out with Nitrosomonas europaea. Keeping in mind that the ammonia oxidizers encompass five different genera

affiliated to two proteobacterial subclasses (section > ‘‘Phylogeny of Ammonia Oxidizers’’), additional species should be investigated to obtain a more encompassing picture of the biochemistry of the ammonia-oxidizing system (Giannakis et al. 1985).

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

108

5000 nitrite ammonia cellnumber

4500 4000 3500 3000

107

2500 2000

cells/ml

nitrite/ammonia [mM]

3

1500 1000 500 0 0

20

40

60

80

100

106 120

time [hrs]

. Fig. 3.7 Growth of Nitrosomonas eutropha in the presence of ammonia, 2,315 parts per million (ppm) acetylene, and hydroxylamine (4 mmol) as substrate (48–96 h). The AMO was inhibited by acetylene, while cell growth was detectable after a lag phase of 2 days. Most of the hydroxylamine undergoes deterioration in contact with atmospheric oxygen. As calculated from additional nitrite formation, 400 mmol of hydroxylamine was oxidized to nitrite, resulting in an increase of cell number

Ammonia and Hydroxylamine as Substrates The overall process of ammonia oxidation to nitrite may be characterized as a two-stage process: NH3 þ 0:5O2 ! NH2 OH NH2 OH þ H2 O ! HNO2 þ 4Hþ þ 4e

ð3:11Þ ð3:12Þ

However, this two-stage scenario is a simplification. For lithotrophic ammonia oxidizers, ammonia is essential as the primary substrate. The intermediate hydroxylamine (NH2OH) is the real energy source. The coupling between ammonia and hydroxylamine oxidation, a complex mechanism not yet established in detail, is suggested by several observations. The addition of hydroxylamine to ammonia-oxidizing cells shortened the lag phase of ammonia oxidation (Hooper 1969), probably by providing reductants to the monooxygenase. It is generally assumed that partial reduction of c-type cytochromes is necessary to start ammonia oxidation. Cytochrome reduction was attained by addition of hydroxylamine to cell-free preparations of Nitrosomonas europaea (Suzuki et al. 1981). If both ammonia and hydroxylamine are used, the molar growth yield of hydroxylamine was found to be twice the amount of ammonia (Bo¨ttcher and Koops 1994; De Bruijn et al. 1995). On the other hand, increasing amounts of hydroxylamine are inhibitory to ammonia oxidation (Hyman and Wood 1984b; Poth and Focht 1985; Abeliovich and Vonshak 1993), probably due to imbalancing the redox state of AMO and HAO (Wood 1986). Another result is more difficult to understand. All attempts to grow ammonia oxidizers on hydroxylamine as the only substrate have failed, although hydroxylamine is oxidized to nitrite (Lees 1952; Hoffman and Lees 1953; Engel and Alexander

1958; Nicholas and Jones 1960). This failure is most likely not caused by the toxicity of hydroxylamine, because addition of hydroxylamine in the presence of ammonia promotes substrate oxidation and cell growth. As demonstrated recently, Nitrosomonas eutropha cells are capable of growing on hydroxylamine as the only substrate when AMO is simultaneously inhibited by acetylene (S. Oesterreicher, personal communication; > Fig. 3.7). Without addition of acetylene, N. eutropha cells 3lyse within 3 days when hydroxylamine is oxidized to nitrite, although within the first day NADH and ATP are still formed (C. Look, personal communication). It is important to note that during these experiments, ammonia was present as nitrogen source because hydroxylamine could not be assimilated. The observation that reduction of a functionally active AMO in the absence of ammonia leads to cell death could be explained by the formation of toxic oxygen radicals by this enzyme under these conditions. This suicidal activity of ammonia oxidizers also might cause nitrification breakdown in wastewater treatment plants, if (1) plenty of organic substrate is available as additional alternative electron donor and (2) ammonia is present in very low concentrations.

Enzymes Involved in Ammonia Oxidation Ammonia Monooxygenase The first intermediate of ammonia oxidation is assumed to be hydroxylamine (section > ‘‘Genes Encoding AMO, HAO, and Related Enzymes’’). In the presence of hydrazine (an irreversible inhibitor of hydroxylamine oxidation; Nicholas and Jones 1960; Hynes and Knowles 1978), the production of small quantities of

93

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Oxidation of Inorganic Nitrogen Compounds as an Energy Source

hydroxylamine from ammonia was observed (Hoffman and Lees 1953; Yoshida and Alexander 1964). Using 18O2, it could be demonstrated that more than 92 % of the oxygen in hydroxylamine originates from dioxygen (Dua et al. 1979). The enzyme AMO, catalyzing the conversion of ammonia to hydroxylamine, has not yet been purified as active protein, but Hyman and Wood (1985) were able to identify a membrane-associated 14 C-labeled protein, putatively representing a component of AMO, when whole cells of Nitrosomonas europaea were incubated with [14C]acetylene. The N-terminal amino acid sequence of the [14C]acetylene-labeled protein (AmoA) was determined. Based on this sequence, an oligonucleotide was derived and was used to identify and clone the gene amoA. The AmoA protein is a 31.8 kDa (McTavish et al. 1993), probably containing the active site of AMO (Hyman and Arp 1992), and consists of five transmembrane sequences and one periplasmic loop. In the same operon, a second gene amoB is located adjacent to amoA. From the deduced amino acid sequence, the protein has a molecular weight of 43 kDa (Bergmann and Hooper 1994a) and is characterized by two transmembrane domains and two periplasmic loops (Vanelli et al. 1996). Upstream of the genes amoA and amoB, a third open reading frame amoC is located which might encode a chaperone helping the AmoA and AmoB protein subunits to integrate into the membrane properly (Klotz et al. 1997). Indirect evidence indicates that AMO is a copper-containing monooxygenase (Rees and Nason 1966; Tomlinson et al. 1966; Loveless and Painter 1968; Dua et al. 1979; Hollocher et al. 1981; Wood 1988a; Hooper and Terry 1973). Quantitative immunoblot analysis using polyclonal antibodies revealed that total cell protein of Nitrosomonas eutropha consisted of approximately 6 % AmoA and AmoB, when cells were grown using standard conditions (Pinck et al. 2001). The specific cellular amount of AMO in cells of Nitrosomonas eutropha was regulated by ammonium concentration. At high ammonium concentrations, less AMO was found than under ammonium-limiting conditions. Furthermore, AMO seems to be strongly protected from degradation. Cells starving 1 year for ammonia still contained high amounts of AMO, although they showed far less ammonia oxidation activity than growing cells. Hence, the amount of AMO does not directly correlate with the activity of ammonia oxidation. Most information about the reactions catalyzed by AMO originates from studies with intact cells. In addition to oxidizing ammonia, AMO can hydroxylate non-growthsupporting substrates such as hydrocarbons and alcohols (Hooper and Terry 1973; Suzuki et al. 1976; Tsang and Suzuki 1982; Hyman and Wood 1983, 1984a, 1984b; Hyman et al. 1985; Voysey and Wood 1987). This is not only of theoretical interest but also could be of importance for microbial ecology (Hall 1986). For example, pure cultures of ammonia oxidizers are able to oxidize methane but could not grow on this alternative electron donor (O’Neil and Wilkinson 1977; Hyman and Wood 1983; Jones and Morita 1983). Recent data, however, suggest that at least in the rice rhizosphere, ammonia oxidizers do not significantly contribute to the methane oxidation (Bodelier and Frenzel 1999;

section > ‘‘Co-oxidation and Inhibition of AMO’’). This capability reflects structural and functional homologies between the ammonia and the methane monooxygenase of ammonia oxidizers and methanotrophs, respectively (Bedard and Knowles 1989). Since substrates or competitive inhibitors of AMO are apolar, it seems reasonable to assume that its active site is hydrophobic. As suggested by Hooper et al. (1997), the reaction is started by the activation of oxygen rather than the substrate. Oxygen might be activated by reduction with a reduced metal-containing center of the enzyme followed by the release of water to form a reactive oxygen species. This compound may extract an electron from the substrate (hydroxylation of the substrate) or interact with nitric oxide to form the real oxidant nitrogen dioxide/dinitrogen tetroxide (see also > Fig. 3.17).

Hydroxylamine Oxidoreductase The key enzyme of hydroxylamine oxidation, HAO, is a multiheme enzyme, located in the periplasmic space (Olson and Hooper 1983; Hooper et al. 1984; Hooper and DiSpirito 1985; section > ‘‘Genes Encoding AMO, HAO, and Related Enzymes’’). The enzyme complex has a relative molecular weight of 180,315–190,315 and consists of an a3 oligomer closely associated with three heme centers including seven c-type hemes and a novel heme, P-460, per monomer (Arciero and Hooper 1993; Hoppert et al. 1995; Igarashi et al. 1997; Bergmann and Hooper 1994b). The P-460 was found to be a CO-binding heme (Hooper et al. 1978; Lipscomb et al. 1982). According to spectroscopic and chemical investigations, the P-460 iron resides in a hemelike macrocycle, but the presumed porphyrin must have some unusual features (Andersson et al. 1984). In total, HAO constitutes about 40 % of the c-type heme of Nitrosomonas europaea (Hooper et al. 1978). The c-type hemes of HAO can be placed into two classes with different oxidation-reduction midpoint potentials and protein environments, respectively (Lipscomb and Hooper 1982; Prince et al. 1983; Hooper 1984a; Collins et al. 1993; Arciero et al. 1991). A detailed discussion of possible interactions of the described redox centers of the HAO can be found in Hooper (1989). Hydroxylamine is supposed to bind at the HAO near the P-460 center. Electrons are released and transferred to c-hemes (Hooper and Terry 1977; Hooper and Balny 1982; Olson and Hooper 1983). Initially, Hooper and Balny (1982) postulated that HAO catalyzes a two-electron dehydrogenation of hydroxylamine and a subsequent net addition of one oxygen atom from dioxygen. Later, they favored a mechanism in which water was the source of the second oxygen atom of the metabolic final product nitrite (Andersson and Hooper 1983; Hooper 1984). The oxidation of hydroxylamine to nitrite was postulated to be a two-step reaction with enzyme-bound nitroxyl (HNO) as an intermediate (Andersson and Hooper 1983): NH2 OH ! ½HNOŠ þ 2Hþ þ 2e

ð3:13Þ

½HNOŠ þ H2 O ! HNO2 þ 2Hþ þ 2e

ð3:14Þ

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

However, in cell-free extracts of Nitrosomonas europaea, nitric oxide was suggested as another possible intermediate of hydroxylamine oxidation (Hooper and Terry 1979). Experiments with 15N-label showed that nitric oxide was produced by hydroxylamine oxidation and not by nitrite reduction. The authors discussed a mixed-function hydroxylation of nitric oxide to be involved in the oxidation from (HNO) to nitrite, with all intermediates being enzyme bound. Miller and Wood (1983) analyzed CO-binding cytochromes of the b type in Nitrosomonas europaea and discussed their possible function in binding nitric oxide resulting from hydroxylamine oxidation.

Electron Flow and Energy Transduction Electron Flow The first step of ammonia oxidation to nitrite, the conversion to hydroxylamine, is endergonic. Thus, hydroxylamine is the real energy-generating substrate. If all subsequent steps of the hydroxylamine oxidation to nitrite are coupled to electron transport chains, a maximum yield of four electrons can result. The number of electrons passing to the terminal oxidase(s), however, is uncertain because four systems (ammonia monooxygenase, nitrite reductase, cytochrome oxidase, and NADH production) are fed with electrons from the oxidation of hydroxylamine to nitrite (Wood 1986). Electrons from HAO reduce cytochrome c554, a 25-kDa tetraheme protein (Andersson et al. 1986). Because both ammonia- and hydroxylamine oxidation seem to be balanced at a steady state, cytochrome c554 is thought to be the first electron transfer branch point. Two of the four electrons released from the hydroxylamine oxidation must pass to the monooxygenase reaction, the latter two flow to a second branch point, for example, cytochrome c552 and then to one of the terminal oxidases cytochrome aa3 (DiSpirito et al. 1986) or nitrite reductase. Once per 5.7 cycles, two electrons are assumed to enter a reverse electron transfer pathway for NADH production (Wood 1986). Cytochrome c554 is a probable candidate for the proposed central role because it is a two-electron carrier (Arciero et al. 1991; Bergmann et al. 1994). The electron carriers downstream have not been investigated in detail. However, the production of nitric and nitrous oxide by Nitrosomonas europaea and N. eutropha suggested that nitric oxide reductase as well as nitrous oxide reductase might be present (Hooper et al. 1997). Yamanaka and Shinra (1974) postulated the path of electrons from HAO to the terminal oxidase to be: HAO ! cytochrome c554 ! cytochrome c552 ! terminal oxidase. However, several other membrane-bound redox carriers have been identified in Nitrosomonas europaea. The function of a tetraheme c-type cytochrome (Cyt c B) is unknown (Bergmann et al. 1994). The periplasmic diheme cytochrome c peroxidase (Arciero and Hooper 1994) of Nitrosomonas europaea might protect enzymes like HAO, which are easily inactivated by hydrogen peroxide (H2O2; Hooper and Terry 1977). Wood (1986) suggested a more conventional construction of the electron transport chain, involving

3

ubiquinone and membrane-bound b- and c-type cytochromes. Wood (1986) discussed a possible proton motive Q cycle as described by Mitchell (1975). Ubiquinone (Q8 species) and membrane-bound cytochromes of types b and c were identified in Nitrosomonas europaea (Hooper et al. 1972; Tronson et al. 1973; Miller and Wood 1983). In > Fig. 3.8, a model of the electron flow in ammonia oxidizers is depicted.

Energy Transduction ATP synthesis driven by proton motive force occurs in Nitrosomonas (Drozd 1976, 1980; Hollocher et al. 1982; Kumar and Nicholas 1982). This energy transduction is assumed to proceed at the level of hydroxylamine oxidation, but the process is not as yet well understood. A simplified scheme would be a two-proton release per electron pair translocated from hydroxylamine to the electron transport chain via the periplasmic located HAO outside the membrane. In addition, the consumption of two protons in the cytochrome oxidase reaction is probably located on the cytoplasmic site of the membrane. However, the exact amount of ATP gained by oxidation of hydroxylamine is not known, since the total number of electrons fed into the energy-generating respiration chain per mol hydroxylamine oxidized varies, depending upon growth stage and environmental conditions. This variation reflects the fact that the production of NADH by reverse electron flow is not constant and coupled to hydroxylamine oxidation by an unknown mechanism. According to Hollocher et al. (1982), the H+/O ratio depends on the substrate concentration. From their measurements, they extrapolated the maximum values to be 3.4 and 4.4 for ammonia and hydroxylamine oxidation, respectively. In addition, Drozd (1976, 1980) stated the maximum P/O ratio from hydroxylamine oxidation to be only one. Measurements of respiration-driven proton translocation indicated the association of only one proton translocation loop, with the transport of two electrons channeled from hydroxylamine to the terminal oxidase.

NADH Production Aleem (1966) showed that cell-free extracts of Nitrosomonas europaea catalyzed an ATP-dependent NAD(P)+ reduction with hydroxylamine as substrate. The reaction was interpreted as ATP-driven reverse electron flow. This hypothesis is in accordance with the postulate that the transmembrane oxidation-reduction loops of respiration chains are reversible, with the exception of the cytochrome c oxidase loop. However, in vivo, the proton motive force resulting from the hydroxylamine oxidation might perhaps drive the reverse direction of the electron flow directly, without support of ATP as previously demonstrated for the nitrite oxidizers Nitrobacter winogradskyi and Nitrobacter vulgaris (Freitag and Bock 1990).

95

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

O

2

2

N2

N

i

O

O

?

N

CM

-R ed

H2 ?

NO

NO

-R e d

?

96

Hydrogenase

? ? Cy

NO

HN

O

2 -R

2

tb c 1 ?

?

2e−

ed

TCC

Acetal Pyruvat Fumarat

NADH+H+ NA

DH/Q ?

HAD+ +2H+

Cy

H2O 0.5 O2+2H+

ne−

C

yt c 552

C

yt c 554

2e−

Ub

ic h

ta a

3

in o n / C y t b c 1

reverse electron

HADH+H+

transport

HAD+ +2H+ N NO O

HN

4e−

O

O2

?

2e−

3

N2 O

NH 3

4H−

HA

H2O

O

4

2H+ AMO

H2O NO NO N

NH 2

OH

H

2 OH

. Fig. 3.8 Model of the electron flow in ammonia-oxidizing bacteria. The part of the figure dealing with nitric oxide, nitrogen dioxide, and dinitrogen tetroxide is hypothetical (section ‘‘> Novel Aspects’’). CM cytoplasmic membrane, i inside the cell/cytoplasmic space, o outside of the cell/periplasmic space, and TCA the tricarboxylic acid cycle (Figure was kindly provided by I. Schmidt)

Co-oxidation and Inhibition of AMO The ammonia monooxygenase (AMO) is a nonspecific enzyme. Ammonia oxidizers are capable of co-oxidizing a range of hydrocarbons (including methane and even xenobiotics), which raised interest in exploiting these microorganisms for bioremediation (Vanelli et al. 1990). The broad substrate range of AMO also is responsible for inhibition of ammonia oxidizers by a variety of substances (> Table 3.1). During oxidation of acetylene via AMO, reactive intermediates that bind irreversibly to AMO are formed in the presence of oxygen. The same mechanism causes the inhibition of AMO by trichlorethylene.

The acetylene inhibition can be ameliorated by high ammonia concentrations via an unknown mechanism (Hyman and Wood 1985). Competitive inhibitors of AMO are methyl fluorides, dimethyl ether (Voysey and Wood 1987; Miller et al. 1993; Hyman et al. 1994), alkanes, alkenes (Hyman et al. 1988), and aromatic compounds (e.g., aniline; Keener and Arp 1994; Voysey and Wood 1987; Hyman and Wood 1983; Jones and Morita 1983). Carbon monoxide (CO) not only binds irreversibly to cytochromes but also competitively inhibits AMO, the enzyme that oxidizes it to carbon dioxide (Tsang and Suzuki 1982; Erickson et al. 1972). Since copper is a cofactor of AMO (Loveless and Painter 1968;

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

3

. Table 3.1 Inhibitors of ammonia oxidation Inhibitor

Optimum

Allylthiourea

10

KCN

5  10

Na2S

10

6b 6b

4b

NH2NH2

2  10

CO (95 % O2, 5 % CO)

0.05b

mCCPc

10

Dinitrophenol (DNP)

2  10

Methylene blue

10

Methanol

5  10

3b

5b 4b

4b 3b

Substrate: NH3a

Substrate: NH2OHa

18

100

22

83

0

9

16

86

8

100

17

128

27

100

0

100

0

100

Ethanol

0.09

0

100

Acetate

0.1b

91

100

0

100

23

50

b

Light

420 lux

Temperature

15  C

a

Nitrite-producing rates (%) of whole cells using ammonia as substrate are listed. For comparison, the respective rates for hydroxylamine oxidation are shown. Nitrite-producing rate of the untreated control equals 100 % b Concentration in mol per liter c m-Chlorcarbonyl cyanide phenylhydrazone Modified from Hooper and Terry (1973)

Hooper and Terry 1973), metal chelators such as allylthiourea and diethyldithiocarbamate are noncompetitive, reversible inhibitors (Lees 1952). In addition to some of the above-mentioned inhibitors, > Table 3.1 lists other inhibitors of ammonia oxidation that do not directly interact with AMO. Ammonia oxidation is much more strongly inhibited by all listed physical parameters and chemical compounds than is hydroxylamine oxidation.

Denitrification Catalyzed by Ammonia Oxidizers Ammonia-oxidizing bacteria not only catalyze aerobic ammonia oxidation but also show denitrifying activity with nitrite as electron acceptor. For example, small amounts of nitric oxide and nitrous oxide are produced during denitrification with ammonia as electron donor at reduced oxygen concentrations (Hooper 1968; Goreau et al. 1980; Remde and Conrad 1990; Stu¨ven et al. 1992). When using 14NH4+ and 15NO2 , Poth and Focht (1985) demonstrated that nitrous oxide was produced at low oxygen tension by nitrite reduction and not by hydroxylamine oxidation. The reaction is thought to be catalyzed by a periplasmic soluble cytochrome oxidase/nitrite reductase induced at low oxygen partial pressure (Miller and Wood 1982; Miller and Nicholas 1985; DiSpirito et al. 1985). Additionally, the formation of dinitrogen was observed (Poth 1986; Bock et al. 1995), indicating that at least some strains of Nitrosomonas possess a nitrous oxide reductase. However, this enzyme has not been isolated as yet from denitrifying ammonia oxidizers.

Ammonia oxidizers show relatively high denitrification activities when they are cultivated under oxygen-limited conditions in the presence of organic matter (mixotrophic growth conditions; Bock et al. 1995). However, under these conditions, ammonia oxidation rates are low (Zart et al. 1996). For this reason, the denitrifying potentials of ammonia oxidizers cannot be efficiently exploited for one-step nitrogen removal in wastewater treatment plants. In the absence of dissolved oxygen, Nitrosomonas eutropha and Nitrosomonas europaea are capable of anoxic denitrification using molecular hydrogen, or simple organic compounds such as acetate, pyruvate, or formate as electron donors and nitrite as electron acceptor (Bock et al. 1995; Abeliovich and Vonshak 1992; Stu¨ven et al. 1992).

Genetics of Ammonia Oxidizers Relatively little information regarding the genetic makeup of ammonia oxidizers is available. Most studies focused on Nitrosomonas europaea (genome of ca. 2.2 Mb) whose genomic sequence is currently being determined [{spider.jgi-psf.org}]. For the other ammonia oxidizers of the b- and l-subclasses of Proteobacteria (section > ‘‘Phylogeny of Ammonia Oxidizers’’), sequence information is restricted to the genes coding for the 16S rRNA (for a review, see Purkhold et al. 2000), the 16S-23S rDNA intergenic spacer region (Aakra et al. 1999), as well as the ammonia monooxygenase operon (Rotthauwe et al. 1995; Purkhold et al. 2000; Alzerreca et al. 1999). Recently, a gene for a copper-containing dissimilatory nitrite reductase (nirK) that has been detected by PCR and was sequenced for several b-subclass ammonia-oxidizing bacteria is under way

97

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Oxidation of Inorganic Nitrogen Compounds as an Energy Source

(Casciotti and Ward 2001). Ammonia oxidizers can also harbor plasmids, as demonstrated by the isolation and characterization of two cryptic plasmids in a Nitrosomonas strain retrieved from activated sludge (Yamagata et al. 1999).

Genes Encoding AMO, HAO, and Related Enzymes Genes coding for enzymes involved in the oxidation of ammonia, particularly the ammonia monooxygenase (AMO), the hydroxylamine oxidoreductase (HAO), and the accompanying cytochromes, have been most intensively studied in N. europaea, which has multiple copies of these primary nitrification genes (section > ‘‘Enzymes Involved in Ammonia Oxidation’’). Nitrosomonas europaea has a duplicated amo operon containing a continuous arrangement of the genes amoC, amoA, and amoB, which are cotranscribed as a 3.5-kb mRNA and encode the three subunits of AMO, AmoC, AmoB, and AmoA (McTavish et al. 1993; Klotz et al. 1997; Sayavedra-Soto et al. 1998). A third copy of amoC, which is not associated with the genes for the other subunits of this enzyme, has recently been identified (SayavedraSoto et al. 1998). Multiple amo operons also have been found in several other ammonia oxidizers (> Table 3.2). Furthermore, N. europaea has at least three copies of each of the genes coding for the hydroxylamine oxidoreductase (hao) and cytochrome c554 (cycA or hcy; McTavish et al. 1993; Hommes et al. 1994). Each copy of the hao gene is located 950 bp upstream of a copy of the hcy gene, but both genes are always found to be within different operons (Bergmann et al. 1994; Sayavedra-Soto et al. 1994). Downstream of two of the hcy genes, an ORF (cycB) predicted to encode another tetraheme cytochrome c was detected (Bergmann et al. 1994). The nucleic acid sequences of the multiple copies of all above-mentioned genes (except for the unlinked amoC genes) are either identical or highly similar within a single ammonia oxidizer species, whereas much lower similarities occur between the respective genes of different

species. Thus, it is likely that the multiple gene copies originated from relatively recent gene duplication events and were not caused by lateral gene transfer (Klotz and Norton 1998). It has been speculated that the presence of multiple genes might (1) allow more-rapid generation of the respective mRNA during ammonia flushes within the local environment of the ammonia oxidizers (Hommes et al. 1998) or (2) be responsible for maintaining a certain ratio of the gene products (Bergmann et al. 1994). In addition to those genes with products directly involved in ammonia oxidation, genes of N. europaea encoding the enolase (eno) and CTP synthase (pyrG) were sequenced (Mahony and Miller 1998). The enolase catalyzes the conversion of 2-phosphoglycerate to phosphoenolpyruvate, and its gene was found to be linked on the chromosome with the pyrG gene, albeit both genes are not cotranscribed. A similar arrangement of both genes is present in the Escherichia coli genome (where they are cotranscribed), though these genes are not linked in other investigated bacterial genomes. Unfortunately, no sequence information regarding the genes involved in CO2 fixation/carboxysome formation of the autotrophic ammonia oxidizers is currently available.

Regulation of AMO and HAO One unusual feature of N. europaea is that it possesses multiple copies of those genes directly involved in ammonia oxidation. This is remarkable, since, with the exception of rRNA and tRNA genes, only relatively few cases of gene duplications have been described for bacteria (e.g., Hass et al. 1992; Sela et al. 1989; Tubulekas and Hughes 1993; Kusian et al. 1995). The significance of the N. europaea ammonia oxidation genes being present in multiple copies has been investigated using techniques for transformation and insertional mutagenesis (Hommes et al. 1996, 1998). Disruption of each of the two amoA copies showed

. Table 3.2 Number of amo operons and amoC copy numbers in different ammonia oxidizers Organism

a

amo operon number

amoC copy numbera

Nitrosomonas europaea ATCC 19178

2

3 3

Nitrosomonas eutropha C-91

2

Nitrosospira briensis C-128

3

4

Nitrosospira sp. NpAV

3

4

Nitrosolobus multiformis ATCC25196

3

4

Nitrosospira sp. 39-19

3

4

Nitrosovibrio tenuis NV-12

2

3

Nitrosococcus oceani

1

1

Nitrosococcus sp. C-113

1

1

Several ammonia oxidizers contain in their genomes an additional amoC copy not linked to other amo genes. In N. europaea, the additional amoC copy has 60 % nucleic acid sequence similarity to each of the other two amoC copies From Norton et al. (1996); Alzerreca et al. (1999), and data from GenBank

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

that each copy was functional in N. europaea and that neither copy is essential in the cell. However, knockout of one of the amoA copies, but not of the other, has a significant influence on the growth rates of the cells (Hommes et al. 1998), suggesting different regulation of each copy. Surprisingly, however, the putative s70-type promotors of both amoA genes were found to be identical (Hommes et al. 2001), indicating that the differential transcription of both genes (Hommes et al. 1998) involves regions upstream of the promotor where the DNA sequences of both copies diverge (Hommes et al. 2001). Similar results were obtained with cells carrying single mutations in each of the amoB genes (Stein et al. 2000). Insertional mutagenesis of each of the three hao gene copies, all of which possess s70-type promotors (Hommes et al. 2001), showed that none of them was essential and that their inactivation could be compensated fully by the two remaining hao genes (Hommes et al. 1996). However, owing to the presence of three hao gene copies, differences in their regulation might only become apparent after simultaneous inactivation of two of the copies. Ammonia-oxidizing bacteria thrive in environments where ammonia is often present in very low concentrations. In these habitats, the capability to efficiently make use of temporal flushes of ammonia might represent an important selective advantage for an ammonia oxidizer. Therefore, the genetic and physiological responses of ammonia oxidizers under conditions of ammonium limitation (ammonium present in amounts that can be metabolized to completion), of starvation (absence of ammonium), and in the presence of excess ammonium were intensively investigated. Nitrosomonas cryotolerans and N. eutropha survive ammonia starvation for at least 25 weeks (Jones and Morita 1985) and 1 year, respectively (Pinck et al. 2001). In contrast to energy-starved heterotrophic bacteria, N. cryotolerans cells after 10 weeks of starvation (1) do not miniaturize, (2) maintain stable levels of intracellular ATP, and (3) show no changes in the total protein, DNA, or RNA levels (Johnstone and Jones 1988). Furthermore, quantitative FISH demonstrated that ammonia oxidizers in activated sludge maintain relatively stable cellular rRNA concentrations during starvation for 1 month or inhibition with allylthiourea for several days (Wagner et al. 1995; Morgenroth et al. 2000). During prolonged starvation for several months or years, ammonia oxidizers lose ammonia-oxidizing activity but still contain significant amounts of AMO inasmuch as this enzyme is degraded more slowly in comparison to the mean cellular protein (Pinck et al. 2001). Under conditions of ammonia starvation, the mRNA of the amo gene disappears within 8 h, though the ammonia and hydroxylamine oxidation activities do not change over a period of 24 h (Stein and Arp 1998a). Limiting ammonium concentrations results in a large loss of ammoniaoxidizing activity (85 %) after 24 h, but it neither affects the steady-state levels of amoA mRNA nor the result in degradation of the AmoA subunit (Stein and Arp 1998a). Interestingly, short-chain alkanes and other substrates having a high binding affinity for AMO ameliorate the inactivating effects of ammonia limitation by protecting the energy-generating activity of

3

N. europaea from potentially toxic by-products of its metabolism (Stein and Arp 1998a, b; section > ‘‘Ammonia and Hydroxylamine as Substrates’’). Interestingly, N. europaea cells grown in biofilms recover much faster after ammonium starvation than their planktonic counterparts. Preliminary data suggest that this phenomenon might be caused by cell-to-cell communication via N-(3-oxohexanoyl)-L-homoserine lactone (Batchelor et al. 1997). As expected, ammonium/ammonia induces the transcription of the ammonia monooxygenase and hydroxylamine oxidoreductase genes as well as the transcription of several additional genes that were not further characterized in Nitrosomonas europaea (Sayavedra-Soto et al. 1996). Furthermore, the activity of AMO is regulated by the presence of ammonia at translational (Hyman and Arp 1995; Stein et al. 1997) and posttranslational (Stein et al. 1997) levels.

Biochemistry of Nitrite-Oxidizing Bacteria The second step of nitrification, the oxidation of nitrite to nitrate, is performed by nitrite-oxidizing bacteria. Although at least four different genera of nitrite oxidizers exist in nature (section > ‘‘Phylogeny of Nitrite Oxidizers’’), most of our knowledge on the physiology and biochemistry of these organisms stems from research on Nitrobacter species and thus cannot be generalized for all nitrite oxidizers. The key enzyme of nitrite-oxidizing bacteria is the membrane-bound nitrite oxidoreductase (Tanaka et al. 1983), which oxidizes nitrite with water as the source of oxygen to form nitrate (Aleem et al. 1965). The electrons released from this reaction are transferred via a- and c-type cytochromes to a cytochrome oxidase of the aa3 type. However, the mechanism of energy conservation in nitrite oxidizers is still unclear. Neither Hollocher et al. (1982) nor Sone et al. (1983) were able to find an electron transport chain linked to proton translocation in nitrite-oxidizing cells of Nitrobacter winogradskyi. The first product of energy conservation was shown to be NADH and not ATP (Sundermeyer and Bock 1981). Except for Nitrobacter, all other isolated nitrite oxidizers are obligate lithotrophs with nitrite serving as the only energy source. Although many strains of Nitrobacter are able to grow heterotrophically, growth is very inefficient and slow (Smith and Hoare 1968; Bock 1976). Additionally, inorganic substrates other than nitrite, namely, nitric oxide, can be used for lithotrophic growth, indicating metabolic diversity among Nitrobacter species (Freitag et al. 1987). In anoxic environments, Nitrobacter cells are able to grow by denitrification (Freitag et al. 1987; Bock et al. 1988). Nitrate can be used as acceptor for electrons derived from organic compounds to promote anaerobic growth. Since the oxidation of nitrite is a reversible process, the nitrite oxidoreductase can reduce nitrate to nitrite in the absence of oxygen (SundermeyerKlinger et al. 1984). Furthermore, the nitrite oxidoreductase copurifies with a nitrite reductase that reduces nitrite to nitric oxide (Ahlers et al. 1990).

99

100

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

Nitrite as a Substrate The utilization of nitrite as an energy source has been the subject of several reviews (Wood 1986; Hooper 1989; Bock et al. 1991; Yamanaka et al. 1981; Tanaka et al. 1983; Sundermeyer-Klinger et al. 1984; Fukuoka et al. 1987). Nitrite is oxidized to nitrate, and the oxygen atom in the nitrate molecule is derived from water (Aleem 1965; Kumar et al. 1983; Hollocher 1984) according to > Eq. 3.15. NO2 þ H2 O ! NO3 þ 2Hþ þ 2e

ð3:15Þ

The two electrons released are transported to oxygen, as described in > Eq. 3.16. 2Hþ þ 2e þ 0:5O2 ! H2 O

ð3:16Þ

The produced nitrate is inhibitory for Nitrobacter species at concentrations between 30 and 65 mM, probably owing to feedback inhibition. The electron flux from nitrite to oxygen could pass the following electron carriers (Bock and Koops 1992): nitrite ! molybdopterin ! iron-sulfur clusters ! cytochrome a1 ! cytochrome c ! cytochrome aa3 ! dioxygen

ð3:17Þ

Enzymes Involved in Nitrite Oxidation Nitrite Oxidoreductase Nitrite oxidation is a reversible process. The enzyme nitrite oxidoreductase (NO2-OR) catalyzes the oxidation of nitrite to nitrate and the reduction of nitrate to nitrite (section > ‘‘Genetics of Nitrite Oxidizers’’). The NO2-OR is an inducible membrane protein present in Nitrobacter cells, which are either grown lithotrophically with nitrite or heterotrophically in the presence of nitrate. Depending upon the enzyme isolation technique, the molecular features of NO2-OR vary considerably. Cytochromes of the a and c type were present when the enzyme of Nitrobacter winogradskyi was solubilized with Triton X-100 and purified by ion exchange and size exclusion chromatography (Tanaka et al. 1983). The purified protein was composed of three subunits of 55, 29, and 19 kDa. Cytochromes a1 and c were also found when n-octylglycoside was chosen as detergent. However, using sodium deoxycholate and subsequent isolation by sucrose gradient centrifugation, only cytochrome c could be detected (Sundermeyer-Klinger et al. 1984). In this preparation, the holoenzyme of Nitrobacter hamburgensis consisted of three subunits with relative weights of 116–130, 65, and 32 kDa. No cytochromes were found when the NO2-OR was isolated from membranes by heat treatment. In this case, only two subunits of 115–130 and 65 kDa were present for Nitrobacter winogradskyi, Nitrobacter vulgaris, and for Nitrobacter hamburgensis (Bock et al. 1990). All preparations of the NO2-OR contain molybdenum (Mo) and iron-sulfur clusters. In membranes of Nitrobacter

winogradskyi, signals attributed to molybdenum were detected by electron proton resonance spectroscopy (Ingledew and Halling 1976). In isolated NO2-OR, molybdenum occurred in the form of molybdopterin (Kru¨ger et al. 1987). The molybdenum content varied between 0.13 (SundermeyerKlinger et al. 1984) and 1.4 g-atoms per molecule (Fukuoka et al. 1987). This difference can probably be explained by the fact that molybdenum often is lost during the enzyme isolation procedure. Molybdenum is essential for nitrite oxidation, and when it is replaced by tungsten, lithoautotrophically growing cells of Nitrobacter hamburgensis are inhibited, whereas heterotrophically growing cells are not. Flavoproteins are absent in NO2-OR preparations. When isolated with Triton X-100, manganese was found to be associated with the NO2-OR (Tanaka et al. 1983). The pH optimum of the NO2-OR for nitrite oxidation differs from that for nitrate reduction. Optimal pH for nitrite oxidation with ferricytochrome c550, ferricyanide, or chlorate as oxidants is about 8.0. With reduced methyl or benzyl viologen as reductants, the optimal pH for nitrate reduction ranges from 6.0 to 7.0. The apparent Km value for nitrite oxidized by the NO2-OR with the aid of different electron acceptors varied with the test conditions between 0.5 and 2.6 mM (Tanaka et al. 1983) or 0.5 and 3.6 mM (Sundermeyer-Klinger et al. 1984), whereas the Km value for nitrate amounted to about 0.9 mM. It is important to note that the specific activities of NO2-OR are influenced by the purification steps of the isolation procedure. As shown in > Table 3.3, the nitrite oxidation activity and the nitrate reduction activity are highest in the membrane fraction. Both activities decrease to about 80 % when NO2-OR is isolated from membranes without detergent. If Triton X-100 or sodium deoxycholate is used for isolation, this effect is even more pronounced (Yamanaka and Fukumori 1988; Sundermeyer-Klinger et al. 1984).

Cytochrome c Oxidase In Nitrobacter species, absorption peaks at 605 nm in difference spectra indicate a cytochrome c oxidase of the aa3 type. This membrane-bound enzyme was purified to an electrophoretically homogeneous state (Yamanaka et al. 1981; Sewell et al. 1972), and the function of cytochrome aa3 was determined as a terminal oxidase by photoactivation of CO-inhibited oxygen consumption. In contrast to mitochondrial terminal oxidases, cytochrome aa3 of Nitrobacter winogradskyi is composed of two subunits with 40 and 27 kDa in a molar ratio of l:l (Yamanaka et al. 1979). One molecule of the enzyme contains two molecules of heme a, two atoms of copper, one atom of magnesium, but no zinc (Yamanaka and Fukumori 1988). The Km values were estimated to be 110 and 24 mM for horse heart cytochrome c and ferricytochrome c (both of which can serve as electron donors) of Nitrobacter winogradskyi, respectively. Phospholipids isolated from Nitrobacter winogradskyi did not stimulate the oxidation rate of native ferrocytochrome c or horse heart cytochrome c (Yamanaka

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

3

. Table 3.3 Activity variations of the nitrite oxidoreductase isolated from mixotrophically grown cells of Nitrobacter hamburgensis depending on the isolation procedure Specific activity (units) Fraction

NO2 -oxidizing activitya

NO3 -reducing activityb

Crude extract, 8,000  g

1.728

2.101

Supernatant

2.338

1.839

Membranes

6.047

3.270

Membranes after heat treatment

2.582

4.882

Purified enzyme

2.506

1.740

a

The unit of activity for NO2 oxidizing was determined with ClO3 as electron acceptor. One unit is defined as the oxidation of 1 mM nitrite per minute and per milligram of protein b The nitrate reductase activity was measured with reduced methyl viologen (MVH) as electron donor. One unit is defined as the reduction of 1 mM nitrate and per minute and per milligram of protein

and Fukumori 1988). If cytochrome aa3 was incorporated in phospholipid vesicles or membrane vesicles, respiratory control was observed, but proton-pumping activity was not (Sone et al. 1983; Sone 1986).

O

CM

i NO2– + H2O

Cyt a1c1

NO2–OR

NO3– + 2H+

Nitrite Reductase In Nitrobacter vulgaris, a membrane-bound nitrate reductase (NiR) was copurified with the nitrite oxidoreductase (Ahlers et al. 1990). The NiR reduces nitrite to nitric oxide, which is released under reduced oxygen partial pressure from the cells to the environment. Therefore, this enzyme seems to be a dissimilatory nitrite reductase of the denitrification type. In the sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) of NO2-OR and NiR, three bands are visible. In addition to the two proteins with Mr 115,000 and 65,000, which are constituents of the NO2-OR, a third protein with Mr 63,000, possibly representing the NiR, is detectable. The pH optimum of the NiR was shown to be 6.1, and the Km value for nitrite was 0.263 mM. The isoelectric point (IEP) was calculated to be at pH 5.5–6.0. Reduced horse heart cytochrome c can serve as an electron donor for nitrite reduction in Nitrobacter winogradskyi and Nitrobacter vulgaris. The biological function of NiR is difficult to understand, since ATP generation has not been detected during nitrite reduction (Freitag and Bock 1990).

Cyt c

Cyt o

2e−

H2O 0,5 O2 + 2H+ Acetate

Cyt bc1 TCC HADH + H+ HADH / Q

HAD+ + 2H+

Cyt a1c1

NO2–OR

Cyt a1c1

NO2–OR

HADH + H+ HAD+ + 2H+ NO2– + H2O

2e− Cyt c

NO3– + 2H+ Cyt aa3

H2O 0,5 O2 + 2H+

nitrite /nitrate

Electron Flow and Energy Transduction As shown in > Fig. 3.9, the first step, the electron transfer from nitrite to cytochrome a1 is catalyzed by the enzyme nitrite oxidoreductase. Cytochrome a1 was shown to be necessary to channel electrons from nitrite to cytochrome c (Yamanaka and Fukumori 1988). Cytochrome a1 of Nitrobacter winogradskyi is not autoxidizable (Tanaka et al. 1983) and shows a typical absorption maximum at 589 nm. It is always found in nitriteoxidizing and nitrate-reducing cells of all Nitrobacter species.

antiport

. Fig. 3.9 Model of the electron flow in Nitrobacter. Depicted are the pathways of denitrification and heterotrophic growth (upper part) and the nitrification pathway (lower part). CM cytoplasmic membrane, i inside the cell/cytoplasmic space, o outside of the cell/periplasmic space, NO2-OR nitrite oxidoreductase, and TCA the tricarboxylic acid cycle (Figure was kindly provided by I. Schmidt)

101

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Oxidation of Inorganic Nitrogen Compounds as an Energy Source

On the other hand, Nitrospira marina does not possess any cytochrome of the a type (Watson et al. 1986). The electrons enter the underlying respiratory chain at the level of cytochrome c (Aleem 1968; Cobley 1976b; Aleem and Sewell 1981; Sundermeyer and Bock 1981; Tanaka et al. 1983). The reduction of cytochrome c is a thermodynamically unfavorable step, which is slow in cell-free extracts. Electrons derived from the nitrite/nitrate couple have a redox potential of Em,7 = +420 mV, whereas those of ferrocytochrome c/ferricytochrome c have a potential of Em,7 = +260 mV. A relatively high nitrite concentration would cause a lowering of the redox potential, but in natural habitats, high nitrite concentrations are rarely found (Schmidt 1982). Actually, a highly active cytochrome aa3 pushes nitrite oxidation by the removal of electrons from cytochrome c. In addition, the concentration of cytochrome aa3 also varies dependent upon the oxygen concentration. SDS-PAGE experiments demonstrated that cells of Nitrobacter vulgaris grown under high oxygen partial pressure possess high nitrite-oxidizing activity and a high cytochrome aa3 content, whereas those cells grown under low oxygen tension have a low activity and a low cytochrome aa3 content (E. Bock, unpublished observation). The nitrite-oxidizing system of Nitrobacter vulgaris can be remodeled by reassociation of n-octylglycoside-isolated NO2OR with cytochrome aa3. The activity of the nitrite-oxidizing system increased with increasing amounts of cytochrome c oxidase (> Fig. 3.10). Present alone, NO2-OR or cytochrome aa3 was unable to oxidize nitrite to nitrate. The in vitro modeling of the nitrite-oxidizing system of Nitrobacter vulgaris shows clearly that both enzymes are essential for the oxidation of nitrite to nitrate, with oxygen as the terminal electron acceptor. At a fixed NO2-OR content, the enzyme activity is regulated by the concentration of cytochrome aa3.

In addition to oxygen, CO2 can serve as electron sink in Nitrobacter. Cytochrome c oxidation generates energy that is necessary for autotrophic carbon dioxide fixation. Since lithoautotrophic growth is inefficient, 85–115 mol of nitrite has to be oxidized to assimilate 1 mol of carbon dioxide (Bo¨meke 1954).

Electron Flow of the Conventional Respiratory Chain The electrons from nitrite meet the underlying respiratory chain at the level of cytochrome c (not shown in > Fig. 3.9). This chain functions in lithotrophically, mixotrophically, and heterotrophically growing cells as well as in endogenous respiring cells of Nitrobacter in the absence or presence of oxygen (> Fig. 3.11). Electrons from NADH + H+ (NADH) pass via flavin mononucleotide (FMN) and ubiquinone to a cytochrome bc1 complex and finally to the terminal oxidase. The responsible NADH oxidase has not yet been isolated. However, FMN was found to be present in heterotrophically grown Nitrobacter cells (Kirstein et al. 1986). Ubiquinone Q10 was the isoprenoid in the respiratory chain (Aleem and Sewell 1984). The cytochrome bc1 is supposed to consist of a cytochrome b560 (55 kDa) and cytochrome c1-550 (32 kDa; M. Rudert, unpublished observations). The terminal oxidase (presented in > Fig. 3.11) is a protein complex isolated from Nitrobacter

NADH + H+

FMN

Specific activity [nmol / mg × min]

102

8 Q

6 55,000

Cyt.b - 560

32,000

Cyt.c1 - 550

25,500

Cyt.c - 552

24,500

Cyt.o - 560

4

2

0 0 Cytochrome

1 c oxidase

2 concentration

3 [mg protein]

. Fig. 3.10 Increase of the specific nitrite-oxidizing activity in cell-free enzyme preparations of Nitrobacter vulgaris. Isolated nitrite oxidoreductase was complemented with increasing amounts of cytochrome oxidase (aa3) for 20 h at 28  C. The specific activity was measured as nitrite oxidized to nitrate with oxygen as the electron acceptor

0,5

O2 + 2H+

H2O

. Fig. 3.11 Components of the conventional respiratory chain of Nitrobacter. The blocks symbolize proteins and their subunits in Mr

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

vulgaris cells (M. Rudert, unpublished observations) consisting of a 25-kDa cytochrome c552 and a 24.5-kDa cytochrome o. This oxidase is assumed to be active at high oxygen partial pressure and is also present in lithotrophically grown cells of Nitrobacter winogradskyi (Aleem and Sewell 1984) as well as in heterotrophically grown cells of Nitrobacter hamburgensis (Kirstein et al. 1986). At low oxygen tension, cytochrome aa3 might perform this function (> Fig. 3.9). Additional electron carriers have been described by different authors. Kurokawa et al. (1987) isolated a flavin adenine dinucleotide-containing flavoenzyme from lithotrophically grown cells of Nitrobacter winogradskyi with NAD(P)H cytochrome c reductase and transhydrogenase activities. Two cytochromes of the b type, b560 and b564, were found in Nitrobacter hamburgensis. Cytochrome b560 is typical for heterotrophically grown cells and might belong to the bc1 complex (Kirstein et al. 1986). The function of cytochrome b564 is unknown. Several additional membrane-bound and soluble cytochromes of the c type have been described (Chaudhry et al. 1981; Miller and Wood 1982). As reported by Tanaka et al. (1982) and Yamanaka et al. (1982), the amino acid composition of a soluble cytochrome c of Nitrobacter winogradskyi is similar to the mitochondrial cytochrome c.

ATP Production A generally accepted concept for the mechanism of energy generation derived from the described electron flow system is not available. Cobley (1976a) reported proton release into the cytoplasm, whereas Wetzstein and Ferguson (1985) detected proton extrusion into the periplasmic space coupled to oxidation of nitrite with artificial electron donors. However, protonpumping activity able to drive a membrane-bound ATPase could neither be measured for nitrite-oxidizing cells nor for nitrite-oxidizing vesicles (Hollocher et al. 1982). Apart from the oxidation of exogenous organic substrates, Nitrobacter cells can oxidize endogenous material, for example, poly-b-hydroxybutyrate; this metabolic activity is called ‘‘endogenous respiration.’’ It has been shown that the oxidation of both exogenic and endogenic matter causes electron flow via a ‘‘normal’’ respiratory chain. Thus, nitrite-oxidizing Nitrobacter can be considered as a regulatory specialist because nitrite oxidation interferes with normal respiration, for example, nitrite oxidation might inhibit endogenous respiration (Eigener and Bock 1975). Changing from endogenous respiration to nitrite oxidation, active cells increased their ATP pool to a maximum of 1 mol of ATP by the oxidation of 1 mol of nitrite (Aleem 1968). All attempts to reproduce this result have failed, but in whole cells and in membrane vesicles, ATP was formed at the expense of NADH oxidation with nitrate (Kiesow 1964; Freitag and Bock 1990) and/or oxygen as electron acceptor (Sewell and Aleem 1979). In Nitrobacter, phosphorylation of ADP is carried out by a membrane-bound ATP synthase. Isolated Nitrobacter ATPase is similar to the F1-ATPase from a thermophilic bacterium (Yamanaka and Fukumori 1988). With respect to ATP

production, Nitrobacter might be best described as a ‘‘normal’’ respiring organism, but this does not explain why heterotrophic growth is so slow.

NADH Production and Cell Growth Lithotrophically grown cells of Nitrobacter winogradskyi and Nitrobacter vulgaris possess an average poly-b-hydroxybutyrate (PHB) content of 10–30 % of the cell dry weight (E. Bock, unpublished observations). This relatively high content indirectly indicates overproduction of NADH. Kiesow (1964) demonstrated in vivo NADH synthesis in nitrite-oxidizing cells by measuring the increase in extinction at 340 nm. Repeating these experiments, we also found NADH formation but only at low oxygen partial pressure. The reaction was sensitive to the uncoupler 2,4-dinitrophenol and insensitive to the ATPase inhibitor N,N0 dicyclohexylcarbodiimide (DCCD; Freitag and Bock 1990). In > Fig. 3.12, the classical scheme of reverse electron flow for generation of NADH is shown. The functional models proposed by Wood (1986) and Hooper (1989) leave many questions unanswered. For example, the authors cannot explain why nitrite-oxidizing cells or spheroplasts of Nitrobacter winogradskyi do not produce a proton gradient, which is necessary to understand reverse electron flow (Hollocher et al. 1982). As stated above, nitrite-oxidizing bacteria are able to grow with nitrite, although the electron transfer from nitrite to cytochrome c is more electronegative than the nitrite/nitrate couple (Aleem 1968; Ferguson 1982). In spite of the existing unfavorable redox potential of cytochrome c, the electrons released from nitrite are promptly removed by cytochrome aa3, so that nitrite oxidation can proceed without energy consumption (O’Kelley et al. 1970). As shown by Cobley (Cobley 1976a, b), the membrane potential has a stimulatory effect on the rate of nitrite oxidation. In experiments with whole cells and membrane vesicles, the nitrite-oxidizing activity decreased in the presence of uncouplers, which collapsed the membrane potential. Thus, even the loss of activity of the isolated NO2-OR might be caused by the loss of the transmembrane electric field, which mediates a conformation

–400

0

+400 +420

mV

NAD+ /NADH

reverse, ATP-consuming electron-transport (+146 kl/mol)

cyt c cyt a1

HNO2 + H2O

HNO3 + 2H+ + 2e–

ATP-generating electron-transport (–74 kl/mol) cyt aa1

+800

0.5O2 + 2H+ + 2e–

H2O

. Fig. 3.12 Classical scheme of ATP-dependent NADH synthesis in nitriteoxidizing cells of Nitrobacter

103

104

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

A

A 340 nm

340 nm

0.03

0.03 b

b 0.02

0.02

0.01

0.01

a 0.00

1

2

3

4

5

6

7

min a

0.00 1

2

3

4

5

6

7

min

–0.01 . Fig. 3.13 NADH formation in whole cells of Nitrobacter winogradskyi in the presence of nitrite (a) and nitric oxide (b) under oxic conditions. NADH production was measured as the increase in absorption at 340 nm

. Fig. 3.14 NADH formation in whole cells of Nitrobacter winogradskyi in the presence of nitrite (a) and nitric oxide (b) under anoxic conditions. NADH production was measured as the increase in absorption at 340 nm

transition between an ‘‘inactive’’ and an ‘‘active’’ form of the enzyme (Tsong and Astumian 1987). Sundermeyer and Bock (1981) were the first to present evidence that NADH synthesis is the primary energy-conserving process in nitrite-oxidizing cells. In addition to nitrite, nitric oxide was shown to be a suitable electron donor for NADH synthesis (Freitag and Bock 1990). > Figures 3.13 and > 3.14 show the formation of NADH (increase in absorption at 340 nm) in whole cells of Nitrobacter winogradskyi. When nitrite was added to aerobic cell suspensions, the dissolved oxygen tension dropped within 5 min to less than 4 % of saturation. As shown in > Fig. 3.13, the NADH pool of the cells first decreased for 5 min and then increased at a constant rate. When nitrite was added to anaerobic cell suspensions, the NADH content increased without any lag phase (> Fig. 3.14). As shown in the figures, the rates of NADH formation with nitric oxide as substrate were faster than those with nitrite. Compared to Em,7 = +420 mV for the nitrite/nitrate couple, the redox potential of the nitric oxide/nitrite couple is Em,7 = +374 mV (Wood 1978), if water is the reactant. Thermodynamically, there is no great difference between the two reactions; nevertheless, nitric oxide was the better substrate than nitrite when Nitrobacter winogradskyi was grown lithoautotrophically. It is generally accepted that NADH generation in Nitrobacter cells is an ATP-independent reaction as shown for Thiobacillus ferrooxidans (Lu and Kelly 1988). But it is still unclear how NADH is synthesized.

Genetics of Nitrite Oxidizers With exception of the 16S rRNA genes (Sorokin et al. 1998; Orso et al. 1994; Teske et al. 1994; Ehrich et al. 1995), no genetic data are available for nitrite-oxidizing bacteria other than Nitrobacter (section > ‘‘Phylogeny of Nitrite Oxidizers’’). For Nitrobacter species, sequence of the 16S-23S rRNA intergenic spacer and partial sequences of the 23S rRNA genes have been determined (Grundmann et al. 2000). Furthermore, the genes encoding the two catalytic core subunits of cytochrome c oxidase of Nitrobacter winogradskyi occur in the same operon. Similar to many ammonia-oxidizing bacteria, N. winogradskyi possesses at least two copies of these genes in its genome (Berben 1996). In addition, the sequences of the Calvin cycle genes were determined for Nitrobacter vulgaris (Strecker et al. 1994) and Nitrobacter winogradskyi (GenBank accession numbers [{http:// www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Search=Nucleotide=AF109915=DocSum}{AF109915}] and [{http://www.ncbi. nlm.nih.gov/entrez/query.fcgi?cmd=Search=Nucleotide=AF10 9914=DocSum}{AF109914}]). In Nitrobacter vulgaris, the genes for the large and small subunits of ribulose-1,5bisphosphate carboxylase/oxygenase, the glyceraldehyde-3phosphate dehydrogenase, and a regulatory protein of the LysR family are located together within one cluster. Another cluster contains the genes for fructose-1,6- and sedoheptulose-1,7biphosphatase, phosphoribulokinase, and fructose-1,6- and sedoheptulose-1,7-bisphosphatealdolase. Furthermore, the

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

genes for both subunits (norA and norB) of the membranebound nitrite oxidoreductase (NO2-OR) from Nitrobacter hamburgensis were sequenced. These genes cluster together with an additional ORF (norX) in the order norA, norX, and norB. The deduced amino acid sequence of protein NorB contains four cysteine clusters with striking homology to those of iron-sulfur centers of bacterial ferredoxins. The b-subunit and the sequenced part of the a-subunit of NO2-OR exhibit significant sequence similarities with the b- and asubunits of the two dissimilatory nitrate reductases of several chemoorganotrophic bacteria including E. coli (Kirstein and Bock 1993). This is consistent with biochemical data that suggest a close functional similarity between both enzyme complexes (Sundermeyer-Klinger et al. 1984; Hochstein and Tomlinson 1988).

Heterotrophic Nitrification The oxidation of ammonia (van Niel et al. 1993), hydroxylamine (Ralt et al. 1981), or organic nitrogen compounds, for example, oximes (Castignetti and Hollocher 1984), to nitrite and nitrate by various chemoorganotrophic microorganisms is called ‘‘heterotrophic nitrification.’’ Heterotrophic nitrification is a cometabolism that is not coupled to energy conservation (Wood 1988b). Thus, growth of all heterotrophic nitrifiers is completely dependent on the oxidation of organic substrates (Focht and Verstraete 1977; Kuenen and Robertson 1987). The final product of heterotrophic nitrification often is nitrite (Castignetti and Gunner 1980), so that heterotrophic nitrification may supply the substrate for lithotrophic nitrite oxidizers and heterotrophic denitrifiers. This additional nitrite production (together with the ability of nitrite oxidizers to grow chemoorganotrophically) might explain why in many environments the number of lithoautotrophic nitrite oxidizers is much higher than that of lithoautotrophic ammonia oxidizers (Kuenen and Robertson 1987). Recently, attention has been driven to heterotrophic nitrifiers because many of them are capable of aerobic denitrification in the presence of organic matter, leading to the complete elimination of dissolved nitrogen compounds with the formation of gaseous nitrogen oxides and/or dinitrogen gas (Castignetti and Hollocher 1984; Robertson et al. 1989; Andersson and Levine 1986; van Niel et al. 1987). Owing to the simultaneous nitrifying and denitrifying activity, nitrification rates of heterotrophic nitrifiers are often underestimated (Castignetti and Hollocher 1984; Kuenen and Robertson 1987). For example, Paracoccus denitrificans (formerly called Thiosphaera pantotropha) produces nitrite from urea, ammonia, and hydroxylamine and is also able to reduce nitrite even under aerobic conditions (Robertson and Kuenen 1983, 1984; Robertson and Kuenen 1988). Therefore, in cultures of this organism, nitrite only accumulates when the nitrite reductase activity is repressed. Biochemically, the ammonia-oxidizing enzyme of Paracoccus denitrificans shows some similarities to the AMO of lithotrophic ammonia oxidizers, for example, the ability to

3

oxidize alkanes, the apparent requirement for copper, and inhibition by light, diethyldithiocarbamate and allylthiourea (Moir et al. 1996a; Crossmann et al. 1997). The purified ammonia-oxidizing enzyme of P. denitrificans contains two polypeptides of 38 and 46 kDa, respectively (Moir et al. 1996a). However, the genes encoding for these polypeptides are not closely related to the amo genes of lithotrophic ammonia oxidizers (Crossmann et al. 1997). The hydroxylamine oxidoreductase from P. denitrificans is a monomeric protein of approximately 18.5 kDa containing nonheme iron (Wehrfritz et al. 1993; Moir et al. 1996b). The environmental importance of heterotrophic nitrifiers is controversial in the literature. Generally, it is assumed that in most environments, the biological conversion of reduced forms of nitrogen to nitrite and nitrate is catalyzed mainly by the lithoautotrophic ammonia- and nitrite-oxidizing bacteria and not by heterotrophic nitrifiers. This reflects that the nitrification rates of heterotrophic nitrifiers are small compared to those of autotrophic nitrifiers (Robertson and Kuenen 1988). Therefore, heterotrophic nitrification was thought to occur preferentially under conditions unfavorable for autotrophic nitrification, for example, in acidic environments (Killham 1986). In such environments, heterotrophic bacteria, fungi, and even some algae might contribute considerably to nitrification (Schimel et al. 1984; Killham 1986, 1987; Robertson and Kuenen 1990; Spiller et al. 1976). But according to recent reports, even in acidic soils, heterotrophic nitrification contributes to overall nitrate production only to a minor extent (Stams et al. 1990; Barraclough and Puri 1995).

Novel Aspects Nitrogenous Oxides Are Essential for Aerobic Ammonia Oxidation Nitrifying as well as denitrifying bacteria contribute to the net production of nitrogenous oxides from soil (Kester et al. 1996, 1997a, b) and from aquatic environments (Xu et al. 1995). This is noteworthy since the gaseous compounds nitric oxide (NO), nitrogen dioxide (NO2), and nitrous oxide (N2O) are of significance for the chemistry of the atmosphere (Johnston 1972; Crutzen 1979; Galbally and Roy 1983). Additionally, nitrous oxide acts as a greenhouse gas (Wang et al. 1976; Andersson and Levine 1986). Furthermore, nitric oxide and to a more moderate extent nitrogen dioxide (Mancinelli and McKay 1983) have strong inhibitory effects on bacteria (Mancinelli and McKay 1983; Shank et al. 1962). Toxicity of nitric oxide is based on its capability to form metal nitrosyl complexes (mainly with heme proteins, iron-sulfur proteins, and copper-containing proteins; Henry et al. 1991), resulting, for example, in the inhibition of cytochrome oxidases (Carr and Ferguson 1990). Furthermore, nitric oxide was shown to form S-nitrosothiols from sulfhydryl groups (Stammler et al. 1992; Hausladen et al. 1996) and to cause C ! T transitions in the DNA (Wink et al. 1991).

105

106

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Oxidation of Inorganic Nitrogen Compounds as an Energy Source

To protect themselves from toxic effects of nitric oxide, many bacteria possess detoxifying enzymes. Denitrifying organisms produce nitric oxide by the activity of the nitrite reductase but are able to keep the internal nitric oxide concentration low by reducing it to nitrous oxide via the NO reductase. Thus, these organisms are strongly dependent on a close functional coupling between both enzymes (Zumft 1993). Consistent with this finding, loss of NO reductase is lethal for denitrifying cells of Pseudomonas stutzeri (Braun and Zumft 1991). Other organisms, for example, Pseudomonas strain PS88, which do not possess the ability to denitrify, are able to detoxify nitric oxide by means of oxidative processes that convert it to nitrate (Baumga¨rtner et al. 1996; Hausladen et al. 1996; Koschorreck et al. 1996). For ammonia-oxidizing bacteria under oxic conditions, NO is also toxic but only in the absence of ammonia (E. Bock, unpublished observation). If ammonia is present, NO and/or NO2 is even essential for aerobic ammonia oxidation. Therefore, the removal of nitric oxide (NO) from cultures of Nitrosomonas eutropha by intensive aeration leads to inhibition of ammonia oxidation. This phenomenon has implications on batch cultivation of ammonia oxidizers in the laboratory (Zart and Bock 1998). Usually, it is necessary to avoid intensive aeration or stirring during the first days of incubation; otherwise, the cells will grow very slowly or even not at all. It was possible to achieve recovery of ammonia oxidation by adding nitric oxide to the air supply. The grade of recovery was dependent on the concentration of nitric oxide supplied (Zart et al. 2000). Inhibition of ammonia oxidation was also observed, when nitric oxide was removed from nitrifying cells of N. eutropha by means of DMPS (2,3-dimercapto-1-propanesulfonic acid) in the presence of Fe3+ ions. The addition of nitric oxide lowered inhibition by DMPS. In another assay, ammonia oxidation of Nitrosomonas eutropha was inhibited by activity of the NO-detoxifying bacterium Pseudomonas PS88 (Baumga¨rtner et al. 1996; Koschorreck et al. 1996) and could be recovered by addition of NO or lowering the activity of the pseudomonad (Zart et al. 2000). The enhancing effect of nitric oxide and nitrogen dioxide on aerobic nitrification and cell growth of Nitrosomonas eutropha could also be demonstrated using fermenter cultures (Zart and Bock 1998). As shown in > Fig. 3.15, the specific activity of ammonia oxidation increased drastically in the presence of nitric oxide and even more if nitrogen dioxide was added instead of nitric oxide. Nitrosomonas eutropha was able to tolerate long exposure (up to 30 days) to nitrogen dioxide or nitric oxide at a concentration as high as 50 ppm when ammonia was oxidized. This is unusual since already 1 ppm nitric oxide is inhibitory for various chemoorganotrophic bacteria (Mancinelli and McKay 1983). This experiment also shows that nitrite is not a potent inhibitor for N. eutropha because nitrite accumulated in this experiment to 100 mM. Thus, ammonia oxidation in N. eutropha was obviously not inhibited by nitrite accumulation as described for N. europaea (Anthonisen et al. 1976; Wullenweber et al. 1978; Drozd 1980). The increase of nitrification rates upon nitrogen dioxide or nitric oxide addition was partly due to a significant increase of cell density. In the presence of 50 ppm of nitrogen dioxide, it was

spec. activity [µmol NH4+ mg protein–1 h–1] 20

15

10

5

0 Without NOX

25 ppm NO

50 ppm NO

25 ppm NO2

50 ppm NO2

. Fig. 3.15 Increase in specific activity of ammonia oxidation in fermenter cultures of Nitrosomonas eutropha upon addition of nitric oxide or nitrogen dioxide to the air supply

possible to obtain up to 2  1010 cells ml 1 of N. eutropha (Zart and Bock 1998), a cell density of ammonia oxidizers, which to our knowledge has never been reported before. Generally, in batch cultures of different Nitrosomonas strains, a cell density of about 2  108 cells ml 1 is rarely exceeded (Engel and Alexander 1958; Prosser 1989). In continuous cultures of N. europaea with complete biomass retention, Tappe et al. (1996) obtained about 5  109 cells ml–1. But it is not clear, so far, how nitric oxide and nitrogen dioxide affect the maximum cell density of these organisms. Beyond this, nitric oxide and nitrogen dioxide had an enhancing effect on the increase of cell number of Nitrosomonas eutropha (Zart and Bock 1998). The specific growth rate (measured as increase in protein) slightly increased upon addition of nitrogen dioxide, but the fission rate increased in a much stronger way. Thus, it is obvious that cell growth was uncoupled from protein increase. Consequently, the cells were depleted of protein when growing in the presence of nitrogen dioxide. This is shown in > Fig. 3.16 where the alteration of cell morphology of Nitrosomonas eutropha grown in presence of 50 ppm of nitrogen dioxide is depicted for a period of 4 weeks. Most striking are the reduction of cell material and the increase of electron-dense inclusion bodies. These inclusion bodies were storage material and resembled glycogen-like particles of Nitrospina gracilis (Watson and Waterbury 1971) and Nitrosolobus multiformis (Watson et al. 1971). Surprisingly, reduction of the protein content per cell was accompanied by increased specific activity of ammonia oxidation. Although nitrification and cell growth of ammonia oxidizers were enhanced by adding nitric oxide or nitrogen dioxide to the air supply of the cultures, the cell yield (cell protein produced per mol of ammonia oxidized) and the energy efficiency slightly decreased. This finding might be due to the aerobic production of N2 induced by the addition of NO2 (Zart and Bock 1998). More than 50 % of the ammonia was oxidized to dinitrogen (N2) and traces of nitrous oxide (N2O) by Nitrosomonas eutropha.

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

c

a

d

3

P

C

ICM

C

P

C

ICM

b

e

0,5 µm

ICM P

P a after inoculation b after 1 week c after 2 weeks d after 3 weeks e after 4 weeks

. Fig. 3.16 Electron micrograph of ultrathin sections of cells of Nitrosomonas eutropha showing the morphological alteration of cells grown in a fermenter aerated with 50 ppm NO2. The cells were harvested from the reactor after inoculation (a) and for 1 (b), 2 (c), 3 (d), and 4 weeks (e) of incubation. ICM intracytoplasmic membranes, C carboxysome, and P unknown particles/electron-dense inclusion bodies

Increasing amounts of supplementary nitrogen dioxide resulted in increasing nitrogen losses (Zart and Bock 1998). Previously, significant nitrogen losses were only obtained with extremely oxygen-limited cultures of ammonia-oxidizing bacteria (Bock et al. 1995; Zart et al. 1996). Recently, it could be demonstrated that the addition of NO2 to cells of Nitrosomonas eutropha grown first under anoxic conditions with hydrogen as electron donor and nitrite as electron acceptor (Schmidt and Bock 1997) and then shifted to oxic conditions significantly reduced the lag for the initiation of ammonia oxidation (Schmidt et al. 2001b). Formation of nitric oxide was up to now interpreted as formation of a by-product of ammonia oxidation without significance for the metabolism of ammonia oxidizers. Inhibition of ammonia oxidation of Nitrosomonas eutropha upon withdrawal of nitric oxide from the culture medium indicates that the production of nitric oxide by ammonia oxidizers seems to be not the formation of a ‘‘waste compound’’ but rather the provision of an important agent for the oxidation of ammonia (Zart et al. 1999). However, nitrogen dioxide rather than nitric oxide could represent the decisive agent, where the latter is acting as precursor for nitrogen dioxide. This hypothesis is based on reports of Schmidt and Bock (Schmidt and Bock 1997, 1998) who described anaerobic oxidation of ammonia by Nitrosomonas eutropha using nitrogen dioxide as an oxygen donor in the AMO reaction. It seems not unlikely that nitrogen dioxide might be involved in the conversion of ammonia to hydroxylamine under oxic conditions as well. In such a case, nitrogen dioxide might act as cosubstrate for the AMO reaction. But since atmospheric nitrogen dioxide concentration hardly exceeds 800 ppb (Galbally and Roy 1983; Baumga¨rtner 1991), the cells would not be able to cover their requirements by consuming it just from the atmosphere. They might rather

produce nitric oxide, which can be oxidized chemically to nitrogen dioxide with dioxygen (Bodenstein 1918). Although the latter reaction proceeds predominantly in the gas phase (Ford et al. 1993; Wink et al. 1993), nitrogen dioxide-consuming reactions (Lewis and Deen 1994) and low concentrations of nitric oxide in the liquid (Pires et al. 1994) can alter the development of the aqueous reaction, so that nitrogen dioxide might be produced in biological systems by oxidation of nitric oxide as it is in the gas phase (Huie 1994). Considering the finding of Dua et al. (1979) that up to 97% of the oxygen of hydroxylamine originates from molecular oxygen, it is important to note that oxidation of ammonia with nitrogen dioxide leads to the formation of hydroxylamine and nitric oxide (Schmidt and Bock 1997). Under oxic conditions, the latter might be reoxidized with dioxygen to form nitrogen dioxide, which would again be available for the oxidation of ammonia by the AMO. Thus, molecular oxygen would not react directly with ammonia but is hypothetically mediated by nitric oxide/nitrogen dioxide. Consequently, it is not necessary to provide nitric oxide or nitrogen dioxide and ammonia in the same ratio since the nitrogenous oxides are permanently recycled. In > Fig. 3.17, a hypothetical model of this NOx cycle is depicted which refers to the three-stage catalytic cycle of the tyrosinase reaction (Shears and Wood 1985). The AMO can have three oxidation states (Shears and Wood 1985; Bedard and Knowles 1989; Keener and Arp 1993). As already mentioned, copper is a constituent of the enzyme, and NO/N2O4 is involved in ammonia oxidation. Therefore, it can be speculated that NO is a cofactor of the AMO. In accordance with Zart and Bock (1998), enzyme-bound N2O4 is the final cosubstrate for oxidizing ammonia to hydroxylamine. Dinitrogen tetroxide (N2O4) is formed from NO by oxidation with molecular oxygen. The AMO consists of three stages: (1) the deoxy form (reduced),

107

108

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

Cu+

Cu+

Anaerobic Ammonia Oxidation Catalyzed by Nitrosomonas eutropha

2 H+

Anaerobic Ammonia Oxidation with Nitrogen Dioxide

N2O4 NH3

Oxy O2

3

1

NH2OH H2O

Cu+

Cu+

Cu2+ Cu2+

2

2 NO

2 NO 2e– Deoxy

Metoxy

. Fig. 3.17 Hypothetical model of an NOx-mediated three-stage cycle driving the oxidation of ammonia to hydroxylamine in ammoniaoxidizing bacteria. This cycle does not contribute to the release of NO during aerobic ammonia oxidation (Stu¨ven and Bock 2001), which is probably caused by chemodenitrification of nitrite at the hydroxylamine oxidoreductase (HAO)

(2) the oxy form (oxidized), and (3) the metoxy form (oxidized). The first stage is oxygen sensitive, and the second and the third one are oxygen stable. The oxygen sensitivity of the deoxy form is caused by an excess of electrons that might lead to the formation of oxygen radicals, which directly or indirectly react with NO, forming peroxy nitrite (OONO). This compound is toxic and destroys the AMO. This working model is in accordance with the observation that purified AMO can oxidize ammonia to hydroxylamine with NADH (H+) as electron donor under anoxic conditions with N2O4 as oxidant. Oxygen was shown to be inhibitory (E. Bock and C. Pinck, unpublished data). Copper in the active center might be responsible for noncovalently binding the nitrogen oxides in form of nitrosyl complexes. As described, oxygen is reacting with NO of the deoxy form (> Fig. 3.17/3), transforming NO to N2O4 (oxy form). The copper ions are oxidized from Cu+ to Cu2+. The N2O4 molecule of the oxy form is the final oxygen donor for ammonia oxidation to hydroxylamine. By this reaction, the enzyme is transferred to the metoxy form (> Fig. 3.17/1), which is subsequently converted by reduction to the deoxy form, thereby completing the cycle (> Fig. 3.17/2). This model also provides an explanation for the inhibition of ammonia oxidation after withdrawal of nitric oxide. As demonstrated recently, acetylene inhibits the aerobic and not the anaerobic ammonia oxidation. Therefore, acetylene is assumed to bind to the deoxy form of the AMO (Schmidt et al. 2001a). Additionally, the oxidation of ammonia with nitrogen dioxide/dinitrogen tetroxide (DG00 = 140 kJ · mol 1) is thermodynamically more favorable than the ‘‘classical’’ oxidation of ammonia with dioxygen (DG00 = 120 kJ·mol 1).

In the absence of dissolved oxygen, ammonia oxidation and cell growth have been observed recently in cultures of Nitrosomonas eutropha (Schmidt and Bock 1997). Those cells were able to replace molecular oxygen by nitrogen dioxide or dinitrogen tetroxide, respectively. Hydroxylamine and nitric oxide were formed in this reaction. While nitric oxide was not further metabolized, hydroxylamine was oxidized to nitrite. However, anoxic ammonia oxidation with nitrogen dioxide was more than tenfold slower than ammonia oxidation with oxygen as electron acceptor. The activity of ammonia oxidizers decreases when the oxygen concentration in the medium decreases too. This can be put down to the fact that the oxidant for ammonia oxidation was limited. Therefore, the organisms need another oxidant for anaerobic ammonia oxidation. For a long time, nitrite was discussed as a suitable oxidant (Broda 1977), but as shown in numerous experiments, it could not serve as electron acceptor for anaerobic ammonia oxidation and cell growth in Nitrosomonas (Bock et al. 1995; Zart et al. 1996).

Anaerobic Ammonia Oxidation in Cell-Free Extracts To reveal the stoichiometry of anaerobic ammonia oxidation, consumption and production of ammonia, nitrogen dioxide, nitric oxide, nitrite, nitrous oxide, and dinitrogen were analyzed in cell-free extracts of N. eutropha. Many attempts were performed to prepare active cell-free extracts of Nitrosomonas europaea under oxic conditions (Suzuki et al. 1970, 1981). One of the most serious problems associated with the characterization of the AMO in extracts has been the instability of the enzyme activity (Suzuki et al. 1974, 1981; Ensign et al. 1993) caused by the sensitivity of reduced AMO to oxygen (C. Pinck and E. Bock, unpublished observation). In contrast, the ammonia-oxidizing enzyme system is stable and active in cell-free extracts under anoxic conditions and thus allowed to characterize the anaerobic ammonia oxidation (Schmidt and Bock 1998). In a helium atmosphere supplied with 25 ppm of nitrogen dioxide, ammonia and nitrogen dioxide were consumed in a ratio of approximately 1:2 by cell-free extracts of Nitrosomonas eutropha. The production of nitric oxide was closely related to the consumption of nitrogen dioxide. Nitric oxide was released in amounts nearly equimolar to the consumption of nitrogen dioxide. The production rate of nitrite was significantly lower than the oxidation rate of ammonia. It is assumed that nitrogen dioxide and nitrite served as acceptors for electrons derived from ammonia oxidation. Approximately 22 % of the nitrite was converted into gaseous nitrogen compounds (nitrogen loss). The main products of denitrification were dinitrogen and traces

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

of nitrous oxide. During the anaerobic ammonia oxidation, hydroxylamine concentrations between 30 and 40 mM were measured. In control experiments with cell-free extracts of Ralstonia eutropha (formerly Alcaligenes eutrophus), Enterobacter aerogenes, and Pseudonocardia nitrificans, neither ammonia oxidation nor nitrogen dioxide consumption could be detected. In sterile control experiments, there was again no ammonia consumption. In addition, no formation of nitric oxide, dinitrogen, or nitrous oxide was measurable. At 25  C and atmospheric pressure, the ratio of nitrogen dioxide (NO2) to its dimer dinitrogen tetroxide (N2O4) is 30:70. To decide whether nitrogen dioxide or dinitrogen tetroxide is the electron acceptor for anaerobic ammonia oxidation, experiments were performed at a temperature of 4  C, conditions under which the dinitrogen tetroxide concentration is almost 100 %. Experiments were performed with crude cell-free extracts in the presence of hydrazine as specific inhibitor for HAO. Although the specific activity of the cell-free extracts decreased significantly at 4  C, anaerobic ammonia oxidation could be detected with a stoichiometry of the converted nitrogen compounds comparable to those observed at 25  C. These results indicate that dinitrogen tetroxide can be used as electron acceptor for anaerobic ammonia oxidation. Based on the observed correlations between ammonia and nitrogen dioxide/dinitrogen tetroxide consumption and nitric oxide and hydroxylamine production, the following equations are proposed. > Equations 3.18 and > 3.19 describe the two half-reactions of the anaerobic ammonia oxidation. The total AMO reaction is presented in > Eq. 3.20: NH3 þ 0:5N2 O4 ! NH2 OH þ NO DG0 0 ¼ þ38 kJ
mol

ð3:18Þ

1

0:5N2 O4 þ 2Hþ þ 2e ! NO þ H2 O DG0 0 ¼

178 kJ
mol

ð3:19Þ

1

NH3 þ N2 O4 þ 2Hþ þ 2e ! NH2 OH þ 2NO þ H2 O 0

DG0 ¼

ð3:20Þ

1

140 kJ
mol

For comparison, the aerobic ammonia oxidation to hydroxylamine is given in > Eqs. 3.21–3.23. NH3 þ 0:5O2 ! NH2 OH DG0 0 ¼ þ17 kJ
mol

ð3:21Þ

1

0:5O2 þ 2Hþ þ 2e ! H2 O DG0 0 ¼

137 kJ
mol

ð3:22Þ

1

NH3 þ O2 þ 2Hþ þ 2e ! NH2 OH þ H2 O DG0 0 ¼

120 kJ
mol

1

ð3:23Þ

The dG00 values of the reactions (> 3.19 and > 3.22) were calculated under the assumption that the reducing equivalents for the AMO are energetically near the ubiquinone level

(+110 mV). > Equations 3.18–3.23 indicate that there are only a few differences between the anaerobic and the aerobic ammonia oxidation of Nitrosomonas eutropha. Instead of molecular oxygen in the course of aerobic ammonia oxidation, dinitrogen tetroxide was used as the electron acceptor, and nitric oxide, an additional product, was released. It appears likely that the same enzyme is responsible for both the aerobic and the anaerobic ammonia oxidation, since (1) hydroxylamine is an intermediate of both reactions, (2) acetylene inhibits the aerobic as well as the anaerobic ammonia oxidation, and (3) anaerobic ammonia oxidation starts immediately after transferring cell-free extracts to anoxic conditions. For the oxidation of hydroxylamine under oxic conditions, no oxygen is needed. Therefore, it can be assumed that under anoxic conditions, hydroxylamine was oxidized according to reaction > 3.24 as well. NH2 OH þ H2 O ! HNO2 þ 4Hþ þ 4e DG0 0 ¼

289 kJ
mol

1

ð3:24Þ

Increasing pool sizes of intracellular ATP and NADH indicated energy conservation in the absence of oxygen but presence of nitrogen dioxide (Schmidt and Bock 1998). Under these conditions, reducing equivalents were also used for the reduction of carbon dioxide, resulting in cell growth and excretion of extracellular organic compounds like glycerol into the medium. The relevance of the anaerobic ammonia oxidation with NO2/N2O4 as oxidant for microbial ecology cannot be assessed at present. The nitrogen dioxide concentration of maximal 400–800 ppb in the atmosphere (Crutzen 1979) should be too low to support anaerobic ammonia oxidation. However, this metabolism might occur in oxygen-limited zones, where locally restricted higher nitrogen dioxide concentrations might exist because of the reaction of molecular oxygen and nitric oxide (Nielsen 1992).

Anaerobic Ammonium Oxidation Catalyzed by Deep Branching Planctomycetes Recently, a novel organism was discovered which is capable of catalyzing the anaerobic oxidation of ammonium, with nitrite as electron acceptor (Strous et al. 1999). This organism, which up to now cannot be obtained in pure culture, was enriched from a denitrifying plant reactor where the anaerobic ammonium oxidation process was observed for the first time (Mulder et al. 1995; van de Graaf et al. 1995). By comparative 16S rDNA sequence analysis, the organism was identified as a novel, deep-branching member of the order Planctomycetales, and the name ‘‘Candidatus Brocadia annamoxidans’’ was proposed (Jetten et al. 2001). The 16S rDNA-based molecular diversity surveys and subsequent fluorescence in situ hybridization analyses of several reactors and wastewater treatment plants with anaerobic ammonia-oxidizing activity demonstrated that at least two different genera that form a monophyletic lineage within the Planctomycetales (> Fig. 3.18) can catalyze

109

110

3

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

Rotating-biological-contactor-clone, (AJ 224943) Antarctic-clone (AF146242) sequencing batch methanogenic reactor clone marine-clone(U70712) sludge-clone Antarctic-clone (AF280847) (AB011309) (AF146252) marine-clone(L10943) deep sea sediment clone(AB015552) Antarctic clone (AF142804) Antarctic clone (AF142940)

Candidatus “Brocadia anammoxidans”

Antarctic clone (AF146239) Antarctic clone (AF146246)

Anammox-enrichment-clone (AJ2508827)

marine sediment clone (AJ241009)

Candidatus “Kuenenia stuttgartiensis”

Antarctic-clone (AF146262)

soil-clone (AJ390448) soil-clone (AJ390447)

Isosphaera

Gemmata soil-clone (AJ390447) to outgroups Planctomyces

Pirellula

0.10

. Fig. 3.18 16S rDNA tree of the Planctomycetales. Anaerobic ammonium oxidizers are labeled green. The scale bar represents 10 % estimated sequence divergence (Modified from Schmid et al. 2001)

this process (Schmid et al. 2000, 2001). In most plants analyzed so far, ‘‘Candidatus Kuenenia stuttgartiensis’’ and not ‘‘Candidatus Brocadia annamoxidans’’ is the most abundant anaerobic ammonium oxidizer (Schmid et al. 2000; Jetten et al. 2001). ‘‘Candidatus Brocadia annamoxidans’’ is a chemolithoautotrophic organism with a very low growth rate (0.003 h 1) and a conspicuous ultrastructure. It contains inside the cytoplasm so-called anammoxosomes, membrane-bounded compartments that make up 30–60% of the cell volume and harbor an unusual hydroxylamine oxidoreductase (Jetten et al. 2001). The metabolic pathway of the so-called ANAMMOX process proposed for ‘‘Candidatus Brocadia annamoxidans’’ differs significantly from the known pathway of aerobic, lithotrophic ammonia oxidizers, since ammonium is supposed to be oxidized with hydroxylamine to form hydrazine (N2H4), which is subsequently oxidized to dinitrogen, the main end product of the ANAMMOX process. The four reducing

equivalents generated in this oxidation step are used for the initial reduction of nitrite to hydroxylamine (van de Graaf et al. 1997). Preliminary data suggest that the unusual hydroxylamine oxidoreductase of ‘‘Candidatus Brocadia annamoxidans’’ (Jetten et al. 2001), which has a smaller molecular mass than the respective enzyme of Nitrosomonas and contains several c-type cytochromes, catalyzes the oxidation of hydrazine to dinitrogen. Anaerobic ammonia oxidation occurs only if the cell density of ‘‘Candidatus Brocadia annamoxidans’’ is higher than 1010–1011 cells ml 1. The reason for this cell density-dependent activity is not clear so far. The following > Eqs. 3.25–3.28 summarize the proposed pathway. The overall reaction resembles a process already proposed in 1977 based on theoretical considerations (Broda 1977): HNO2 þ 4½HŠ ! NH2 OH þ H2 O

ð3:25Þ

NH2 OH þ NH3 ! N2 H4 þ H2 O

ð3:26Þ

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

N2 H4 ! N2 þ 4½HŠ

ð3:27Þ

HNO2 þ NH3 ! N2 þ 2H2 O

ð3:28Þ

From > Eq. 3.28, it is obvious that the overall reaction is balanced. No reducing power is gained which is essential for the fixation of carbon dioxide by autotrophic organisms. To gain these reducing equivalents, the cells are required to oxidize nitrite to nitrate. Therefore, the amount of nitrite consumed is about 20 % higher than one could expect from > Eq. 3.28, and nitrate is additionally formed (van de Graaf et al. 1996): HNO2 þ H2 O þ NAD ! HNO3 þ NADH2

ð3:29Þ

The metabolic activity of ‘‘Candidatus Brocadia annamoxidans’’ is strongly inhibited by oxygen, phosphate, acetylene, or shock loading. In addition, organic electron donors or high concentrations of nitrite are inhibitory (van de Graaf et al. 1996). ‘‘Candidatus Brocadia annamoxidans’’ oxidizes ammonium anaerobically about 50-fold faster than Nitrosomonas eutropha with 25 ppm of nitrogen dioxide in the absence of oxygen (Anaerobic Ammonia Oxidation Catalyzed by Nitrosomonas eutropha). Since ANAMMOX was shown to exhibit rather efficient ammonium elimination rates of up to 3 kg of NH4+ m 3 day 1 (van de Graaf et al. 1996), it is suitable for the treatment of wastewater containing much ammonium and little organic chemical oxygen demand (COD). For such wastewater, it has been calculated that the replacement of the conventional nitrogen elimination steps by ANAMMOX would result in a reduction of the operational costs of up to 90 % (Jetten et al. 2001). However, the presence of anaerobic ammonium oxidizers in addition to ammonium nitrite is required in the wastewater. Therefore, partial conventional nitrification, converting approximately half of the ammonium to nitrite, and ANAMMOX have been combined for efficient nitrogen removal from high-strength organic wastewater (summarized in Jetten et al. 2001). It should, however, be noted that very long lag phases are required for obtaining ANAMMOX activity in such plants (Mulder et al. 1995; Jetten et al. 1997), which might be a disadvantage for applications.

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4 H2-Metabolizing Prokaryotes Edward Schwartz . Johannes Fritsch . Ba¨rbel Friedrich Institut fu¨r Biologie, Mikrobiologie, Humboldt-Universita¨t zu Berlin, Berlin, Germany

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 Ecology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126 The Global H2 Budget . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126 Habitats of H2-Evolving Prokaryotes . . . . . . . . . . . . . . . . . . . 130 Habitats of H2-Consuming Prokaryotes . . . . . . . . . . . . . . . . 130 Syntrophy and Interspecies H2 Transfer . . . . . . . . . . . . . . . . . 131 H2-Based Ecosystems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132 Aquifers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 132 Hydrothermal Vents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Physiology: Varieties of H2 Metabolism . . . . . . . . . . . . . . . . . . . 135 H2-Evolving Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 Anaerobic CO Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 Production of H2 as a Byproduct of N2 Fixation . . . 137 Production of H2 as a Byproduct of Phosphite Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 H2-Consuming Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 Aerobic H2 Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 Acetogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 140 Sulfate and Sulfur Reduction . . . . . . . . . . . . . . . . . . . . . . . . 140 Fe(III) Reduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142 Dehalorespiration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 142 Anoxygenic Photosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . 142 Fumarate Respiration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 Ancillary Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 Classification of Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144 [FeFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144 [Fe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144 Biochemistry of the Hydrogen-Converting Enzymes . . . . . . 144 [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 The Basic Module . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145 The [NiFe] Active Site . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 147 Catalytic Reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 148 Interaction of [NiFe] Hydrogenases with Cytochromes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 149 A Histidine Kinase Is the Target for the HydrogenSensing [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . 150 Multimeric [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . 150 Energy-Converting [NiFe] Hydrogenases . . . . . . . . . . . 151 [FeFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Molecular Characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Catalytic Reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153

Diversity of [FeFe] Hydrogenases and Related Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 154 [Fe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 156 Molecular Characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . . 156 Catalytic Reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 157 H2 Conversion in the Presence of O2 . . . . . . . . . . . . . . . . . . . 157 Biogenesis of Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 Maturation of [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . 159 Membrane Translocation of Hydrogenases . . . . . . . . . . . . . 161 Additional Functions Involved in the Maturation of Periplasmically Oriented Membrane-Bound [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162 Maturation of [FeFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . 163 Maturation of [Fe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . 164 Genetic Organization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164 [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164 Archaeal Membrane-Bound Hydrogenases . . . . . . . . . . . . . 165 Bacterial Membrane-Bound Hydrogenases . . . . . . . . . . . . . 165 The Cytoplasmic F420-Nonreactive Hydrogenases . . . . . . 166 The F420-Reactive Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . 166 The Cytoplasmic, NAD-Reducing Hydrogenases . . . . . . . 167 Multimeric H2-Evolving Hydrogenases . . . . . . . . . . . . . . . . . 168 Putative High-Affinity Hydrogenases . . . . . . . . . . . . . . . . . . . 168 [FeFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 [Fe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 Regulation of Hydrogenase Genes . . . . . . . . . . . . . . . . . . . . . . . . . 169 [NiFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 169 [FeFe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 [Fe] Hydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 Evolutionary Aspects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 Abstract The reversible splitting of H2 into protons and electrons is a key process in the metabolism of many prokaryotes and has been studied extensively in a wide range of bacteria and archaea. Environmental H2 is an energy source for aerobic H2 oxidizers, methanogens, acetogens, and sulfate reducers and is a source of reducing power for anoxygenic phototrophs. H2 is released as a terminal metabolic product of both facultative and obligate fermenters. It is a byproduct of N2 fixation and phosphite oxidation. The H2-consuming and H2-evolving processes of microorganisms impact the global atmospheric H2 balance. N2 fixation in seas and lakes is a significant source of atmospheric

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_65, Springer-Verlag Berlin Heidelberg 2013

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H2. Soils are a major H2 sink. Entirely H2-based microbial ecosystems are widespread on the planet. The most important of them consists of the granitic layers of the planet’s crust, which on aggregate harbor a huge fraction of the total biomass on Earth. More spectacular are the submarine hydrothermal vents spewing H2-rich fluids. Current scenarios of pre- and protobiotic evolution envisage such sites as the cradle of terrestrial life. Based on their metal content, hydrogenases, the enzymes which catalyze the splitting of H2, can be divided into three groups of independent phylogenetic origin: [NiFe], [FeFe], and [Fe] hydrogenases. Three-dimensional structures available for representatives of all three groups reveal some remarkable features of these enzymes. The actual catalyst is a NiFe or Fe metallocomplex encased in a protein. Tunnels in the protein allow H2 to access or egress from the active site. A series of FeS clusters form an electrical circuit connecting the active site with binding sites (for cytochromes, pyridine nucleotides, and other redox partners) at the surface of the enzyme. The assembly and insertion of the active-site metallocomplex into the hydrogenase apoenzyme is an intricate, multistep process requiring several specialized accessory proteins. The genetic determinants for the hydrogenase catalytic components and for the accessory proteins are solitary or clustered. The mechanisms governing the expression of hydrogenase genes vary depending on physiological context. In obligate fermenters, for instance, expression of hydrogenase genes is typically constitutive. In facultative H2 oxidizers, on the other hand, hydrogenase gene expression is controlled by H2-sensing regulatory proteins. The diversity of metabolic processes involving H2 as an intermediate and the ubiquitous occurrence of hydrogenases in microbes testify to the importance of H2 metabolism in primeval cellular life forms.

Introduction During the era of prebiotic evolution, which culminated in the appearance of cellular life forms, the earth had a reducing atmosphere. Fueled by volcanic activity and by magmatic outgassing, levels of atmospheric molecular hydrogen (H2) may have been higher than in the present-day atmosphere. This is the subject of ongoing debate (Tian et al. 2005; Catling 2006) and estimates vary over several orders of magnitude (Walker 1977; Kasting 1993). Various evolutionary scenarios envisage primeval life forms with H2-based metabolism (Stetter 1992; Wa¨chtersha¨user 1992; Edwards 1998). Some of the recent theories postulate H2-rich effluents of hydrothermal vents as the matrix for the evolution of early biochemical systems (Martin et al. 2008; Martin 2012). With the advent of oxygenic photosynthesis (a process which may have originated as early as 3.5 billion years ago), oxygen began accumulating in the atmosphere (Walker 1977; Hayes 1983; Blankenship 1992; Kasting et al. 1992; Nisbet and Fowler 1999). The transition to an oxic atmosphere was accompanied by the rise of O2-respiring organisms, which expanded to habitats all over the Earth’s surface, whereas the strictly H2-dependent organisms were confined to

specialized niches. Nevertheless, H2 continues to be an important and widespread metabolite in both the archaeal and bacterial realms of the microbial world, in oxic as well as in anoxic habitats. The physiological role of H2 in microbes is dual: Firstly, H2 is a growth substrate, that is, a source of energy and reductant. Prokaryotes of different metabolic types, such as methanogens, anoxygenic phototrophs, and aerobic knallgas bacteria, exploit H2. Secondly, H2 production is a means of dispersing excess reductant from fermentative metabolism. Hydrogen is among the fermentation products of both facultative fermenters, such as Escherichia coli, and obligate fermenters such as Clostridium pasteurianum. This chapter surveys the H2-metabolizing prokaryotes. It covers both H2 evolution and H2 consumption in archaea and bacteria. A glance at the list of H2-metabolizing prokaryotes (> Tables 4.1 and > 4.2) will convince the reader that H2 metabolism is not limited to specialized microbes grouped in a few taxonomic units. The comprehensive approach chosen here cuts across the boundaries of the physiological groups described in classical microbiology textbooks. This is appropriate since the physiology of many H2 metabolizers can only be explained in the context of syntrophic associations. The common denominator of the disparate taxonomic and physiological groups treated here is hydrogenase, the enzyme responsible for reversibly catalyzing all or part of the reaction: H2 $ H þ Hþ $ 2Hþ þ 2e

ð4:1Þ

It is now agreed that presently known hydrogenases belong to three groups of independent phylogenetic origin. These groups are defined on the basis of their metal content: [FeFe] hydrogenases, [NiFe] hydrogenases (including [NiFe(Se)] hydrogenases), and [Fe] hydrogenases (formerly referred to as ‘‘metal-free hydrogenases’’). Thus, hydrogenases constitute an example of convergent evolution. The [FeFe] and [NiFe] hydrogenases are particularly interesting in this context. Although the representatives of these two groups are not related, both the architectures of their active sites and the mechanisms of the chemical reaction catalyzed by them reveal striking similarities. The past decades have seen rapid advances in our knowledge of the genetic basis of H2 metabolism on the one hand and of the structure and catalytic mechanism of hydrogenases on the other. Accordingly, the major part of this chapter is devoted to an overview of these genetic, biochemical, and spectroscopic studies. Hydrogen evolution and consumption is not catalyzed by hydrogenases alone. Nitrogenase, the enzyme which catalyzes the production of NH3 from atmospheric dinitrogen, produces H2 as a byproduct of N2 fixation. Nitrogen-fixing prokaryotes are mentioned here because of their important contribution to the global H2 flux and because N2 reduction is an important physiological context of hydrogenase. An in-depth treatment of this subject is, however, beyond the scope of this article. Hydrogenases are also found in eukaryotes. The hydrogenases of the chloroplasts of green algae, for instance, have

H2-Metabolizing Prokaryotes

. Table 4.1 H2-metabolizing bacteria Group

Order

Speciesa

Eb References

Aquificales

Aquifex aeolicus

G

Deckert et al. (1998)

Aquifex pyrophilus

P

Huber et al. (1992)

P

Kryukov et al. (1983)

Hydrogenobacter hydrogenophilusc Hydrogenobaculum acidophilum

Cyanobacteria

Chlorococcales

d

P

Shima and Suzuki (1993)

Hydrogenobacter thermophilus

P

Kawasumi et al. (1984), Shiba et al. (1984)

Hydrogenothermus marinus

P

Sto¨hr et al. (2001b)

Thermocrinus ruber

P

Huber et al. (1998)

Balnearium lithotrophicum

P

Takai et al. (2003a)

Sulfurihydrogenibium subterraneum P

Takai et al. (2003b)

Desulfurobacterium pacificum

P

L’Haridon et al. (2006)

Desulfurobacterium atlanticum

P

L’Haridon et al. (2006)

Persephonella marina

P

Go¨tz et al. (2002)

Persephonella guaymasensis

P

Go¨tz et al. (2002)

Persephonella hydrogenophila

P

Nakagawa et al. (2003)

Thermovibrio ammonificans

P

Vetriani et al. (2004)

Thermovibrio ruber

P

Huber et al. (2002)

Phorcysia thermohydrogiphila

P

Pe´rez-Rodrı´guez et al. (2011)

e

Synechococcus PCC 6301

P

Peschek (1979), Howarth and Codd (1985)

A

Schmitz et al. (1995), Schmitz and Bothe (1996)

Synechococcus PCC 6307

P

Howarth and Codd (1985)

Synechocystis PCC 6803

P

Howarth and Codd (1985)

Synechocystis PCC 6714

P

Howarth and Codd (1985)

Synechocystis PCC 6308

P

Serebryakova et al. (1996)

Cyanothece PCC 7822

P

van der Oost et al. (1989)

Microcystis PCC 7820

P

Howarth and Codd (1985)

f

Nostocales

Microcystis PCC 7806

P

Moezelaar and Stal (1994)

Nostoc sp. PCC 73102

P

Lindblad and Sellstedt (1990)

Nostoc sp.g PCC 7937

PA Mikheeva et al. (1995), Serebryakova et al. (1994) I

Nostoc muscorum PCC 7120

I Anabaena cylindricah PCC 7122

Flexibacteria

Ewart and Smith (1989)

Stigonematales

Fischerella muscicola PCC 73103

A

Lambert and Smith (1980)

Oscillatoriales

Oscillatoria chalybea

P

Bader and Abdel-Basset (1999)

Oscillatoria limosa

P

Heyer et al. (1989)

Microcoleus chthonoplastes

P

Moezelaar et al. (1996)

P

Miroshnichenko et al. (2003d)

Deferribacter desulfuricans

P

Takai et al. (2003a)

Deferribacter autotrophicus

P

Slobodkina et al. (2009b)

Oceanithermus profundus

P

Miroshnichenko et al. (2003b)

Vulcanithermus mediatlanticus

P

Miroshnichenko et al. (2003c)

Chloroflexus aurantiacus

PA Holo and Sirevag (1986)

Deferribacteres Deferribacterales Deferribacter abyssi

DeinococcusThermus

Houchins and Burris (1981b)

PA Bothe et al. (1977), Lambert and Smith (1980) I

i

Serebryakova et al. (1996)

PA Houchins and Burris (1981a)

Thermales

4

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4

H2-Metabolizing Prokaryotes

. Table 4.1 (continued) Speciesa

Group

Order

Firmicutes

Actinomycetales Frankia sp. Bacillales Clostridiales

Eb References A

Sellstedt (1989), Murry and Lopez (1989)

Streptomyces thermoautotrophicus

P

Gadkari et al. (1990)

Bacillus schlegelii

P

Schenk and Aragno (1979)

Bacillus tusciae

P

Bonjour and Aragno (1984)

Clostridium pasteurianum

P

Nakos and Mortenson (1971)

I

Chen and Mortenson (1974), Chen and Blanchard (1978)

P

Gray and Gest (1965)

Clostridium acetobutylicum

I

Vasconcelos et al. (1994)

Clostridium kluyveri

P

Bornstein and Barker (1948)

Clostridium tetanomorphum

P

Woods and Clifton (1938)

Clostridium tepidiprofundi

P

Slobodkina et al. (2008)

Clostridium caminithermale

P

Brisbarre et al. (2003)

Clostridium aceticum

P

Braun et al. (1981), Braun and Gottschalk (1981)

Clostridium mayombei

P

Kane et al. (1991)

Clostridium magnum

P

Bomar et al. (1991)

Clostridium scatologenes

PA Ku¨sel et al. (2000)

Eubacterium limosum

P

Sharak-Genthner and Bryant (1987)

Carboxydothermus hydrogenoformans

P

Svetlichnyi et al. (1991)

I

Soboh et al. (2002)

Ruminococcus albus

P

Miller and Wolin (1973)

Ruminococcus hydrogenotrophicus

P

Bernalier et al. (1996a)

Ruminococcus productus

P

Lorowitz and Bryant (1984), Bernalier et al. (1996b)

Ruminococcus hansenii

P

Bernalier et al. (1996b)

Ruminococcus schinkii

P

Rieu-Lesme et al. (1996)

Syntrophomonas wolfei

PA McInerney et al. (1979), McInerney et al. (1981a)

Megasphaera elsdenii

I

Van Dijk et al. (1979), Atta and Meyer (2000)

Tepidibacter thalassicus

P

Slobodkin et al. (2003)

Caminicella sporogenes

P

Alain et al. (2002)

Thermincola carboxydiphila

P

Sokolova et al. (2005)

Dethiosulfatibacter aminovorans

P

Takii et al. (2007)

Desulfitobacterium dehalogenans

P

Utkin et al. (1994)

Rhodococcus opacus

P

Aggag and Schlegel (1973)

Mycobacterium gordonae

P

Park and DeCicco (1974)

Sporomusa termitida

P

Breznak et al. (1988)

Sporomusa sphaeroides

PA Dobrindt and Blaut (1996)

Sporomusa sylvacetica

P

Kuhner et al. (1997)

Acetonema longum

P

Kane and Brezhnak (1991a)

Desulfotomaculum orientis

P

Klemps et al. (1985)

A

Cypionka and Dilling (1986)

Desulfotomaculum geothermicum

P

Daumas et al. (1988)

Desulfotomaculum thermocisternum

P

Nilsen et al. (1996)

Desulfotomaculum thermosubterraneum

P

Kaksonen et al. (2006)

Acetobacterium woodii

P

Balch et al. (1977), Ljungdahl and Wood (1982)

I

Ragsdale and Ljungdahl (1984)

P

Kotsyurbenko et al. (1995)

Acetobacterium bakii

H2-Metabolizing Prokaryotes

. Table 4.1 (continued) Group

Order

Thermoanaerobacteriales

Speciesa

Eb References

Acetobacterium paludosum

P

Kotsyurbenko et al. (1995)

Acetobacterium fimetarum

P

Kotsyurbenko et al. (1995)

Thermoanaerobacter kivui

PA Leigh et al. (1981), Daniel et al. (1990), Pusheva et al. 1991

Moorella thermoaceticaj

PA Fontaine et al. (1942), Drake (1982), Daniel et al. 1990 P

Thermolithobacterales

Moorella thermoautotrophica

PA Clark et al. (1982), Wiegel et al. (1981)

Thermacetogenium phaeum

P

Hattori et al. (2000)

Carboxydothermus siderophilus

P

Slepova et al. (2009)

Thermovenabulum ferriorganovorum

P

Zavarzina et al. (2002)

Caldanaerobacter subterraneus

P

Sokolova et al. (2001)

Thermolithobacter ferrireducens

P

Sokolova et al. (2007)

Thermolithobacter carboxydivorans

P

Sokolova et al. (2007)

P

Tsai et al. (1995)

Halanaerobiales Halanaerobium alcaliphilum

Proteobacteria

Pezacka and Wood (1984), Martin et al. (1983), Kerby and Zeikus 1983

Acethalobium arabaticum

P

Zhilina and Zavarzin (1990)

Selemonadales

Thermosinus carboxydivorans

P

Sokolova et al. (2004)

Bacteroidales

Acetomicrobium flavidum

P

Soutschek et al. (1984)

I

Mura et al. (1996)

a Class

Renobacter vacuolatum

P

Malik and Schlegel (1981)

Aquaspirillum autotrophicum SA32

P

Aragno and Schlegel (1978)

Bradyrhizobium japonicum

P

Hanus et al. (1979), Emerich et al. (1979)

Paracoccus pantotrophusk

A

McCrae et al. (1978)

I

Harker et al. (1985)

P

Ku¨hnemund (1971)

A

Schneider and Schlegel (1977)

I

Knu¨ttel et al. (1989), Sim and Vignais (1978)

Methylosinus trichosporium OB3b

A

Chen and Yoch (1987)

Rhizobium leguminosarum

P

Dixon (1968), Nelson and Salminen (1982)

Rhodobacter capsulatus

P

Yen and Marrs (1977), Madigan and Gest (1978), Madigan and Gest (1979)

I

Colbeau et al. (1983)

Rhodobacter sphaeroides

P

Uffen and Wolfe (1970)

Rhodospirillum rubrum

P

Ormerod and Gest (1962), Gest (1954), Gorrell and Uffen (1978), Voelskow and Scho¨n (1980), Uffen 1981

I

Adams and Hall (1979a)

Rhodopseudomonas palustris

P

Qadri and Hoare (1968), Uffen and Wolfe (1970)

Thiorhodococcus minus

P

Guyoneaud et al. (1997)

Thiocystis violacea

P

Winogradsky (1888)

Rhodomicrobium vannielii

P

Duchow and Douglas (1949)

Xanthobacter flavus 301

P

Malik and Claus (1979)

Xanthobacter autotrophicus

P

Schneider et al. (1973), Baumgarten et al. (1974)

I

Schink (1982)

A

Eberhardt (1969), Schneider and Schlegel (1977)

P

Meyer and Schlegel (1978)

I

Santiago and Meyer (1997)

Azospirillum lipoferum

P

Malik and Schlegel (1981)

Ancylobacter aquaticus

P

Malik and Schlegel (1981)

Oligotropha carboxidovorans

4

123

124

4

H2-Metabolizing Prokaryotes

. Table 4.1 (continued) Group

Order

Speciesa

Eb References

b Class

Acidovorax facilis

P

Willems et al. (1990)

Acidovorax delafieldii

P

Willems et al. (1990)

Variovorax paradoxus

P

Davis et al. (1970)

A

Schneider and Schlegel (1977)

Achromobacter ruhlandii

P

Packer and Vishniac (1955), Aragno and Schlegel (1977)

Alcaligenes latus H-4

P

Palleroni and Palleroni (1978)

I

Pinkwart et al. (1983)

Alcaligenes hydrogenophilus

P

Ohi et al. (1979)

Hydrogenophaga flava

P

Willems et al. (1989)

Hydrogenophaga pseudoflava

P

Willems et al. (1989)

Hydrogenophaga palleroni

P

Davis et al. (1970)

P

Willems et al. (1989)

Hydrogenophaga taeniospiralis

P

Lalucat et al. (1982), Willems et al. (1989)

Ralstonia eutropha H16

P

Eberhardt (1966)

I

Schneider and Schlegel (1976), Schink and Schlegel (1979), Bernhard et al. (2001)

Ralstonia metallidurans CH34

PA Mergeay et al. (1985)

Hydrogenophilus hirschii

P

Sto¨hr et al. (2001a)

Hydrogenophilus thermoluteolus

P

Hayashi et al. (1999)

P

Wertlieb and Vishniac (1967), Uffen (1976)

Thiobacillus plumbophilus

P

Drobner et al. (1992)

Pseudomonas saccharophila

PA Bone (1960), Bone et al. (1963), Podzuweit et al. 1983

Rubrivivax gelatinosus

g Class

l

Azotobacter chroococcum Azotobacter vinelandii

Lee and Wilson (1943) van der Werf and Yates (1978)

P

Hyndman et al. (1953), Hyndman et al. (1953), Wong and Maier 1985

I

Seefeldt and Arp (1986)

Derxia gummosa

P

Pedrosa et al. (1980)

Acidithiobacillus ferrooxidans

P

Drobner et al. (1990), Fischer et al. (1996)

Escherichia coli

P

Stephenson and Stickland (1931), Peck and Gest (1957)

A

Krasna (1980, 1984)

I

Adams and Hall (1979b), Ballantine and Boxer (1985), Sawers and Boxer (1986)

Salmonella typhimurium

A

Krasna (1980)

Citrobacter freundii

A

Krasna (1980)

Klebsiella oxytoca

I

Wu et al. (2011)

Allochromatium vinosum

I

Gitlitz and Krasna (1975)

Shewanella putrefaciens

P

Lovley et al. (1989)

Thiocapsa roseopersicina BBS

P

Bogorov (1974), Gogotov et al. (1974)

I

Zorin and Gogotov (1975), Gogotov et al. (1976)

P

Nishihara et al. (1990, 1991)

I

Nishihara et al. (1997)

P

Stanley and Dalton (1982)

Hydrogenovibrio marinus Methylococcus capsulatus Bath

d Class

P I

A

Hanczar et al. (2002)

Pseudomonas hydrogenovora

P

Kodama et al. (1975), Igarashi et al. (1980)

Desulfomicrobium norvegicumm

P

Genthner et al. (1997)

Desulfomicrobium baculatumn

P

Rozanova et al. (1988), Lampreia et al. (1991)

Desulfomicrobium apsheronum

P

Rozanova et al. (1988)

Desulfobacterium autotrophicum

P

Brysch et al. (1987)

H2-Metabolizing Prokaryotes

. Table 4.1 (continued) Group

Order

Speciesa Desulfovibrio vulgaris

e Class

Thermodesulfo- Thermodesulfobacteria bacteriales Thermotogae

Thermotogales

Eb References G

Strittmatter et al. (2009)

P

Hatchikian et al. (1976), Badziong et al. (1978), Traore et al. 1983

I

Yagi (1970), van der Westen et al. (1978)

Desulfovibrio fructosovorans

P

Malki et al. (1997)

Desulfovibrio desulfuricans

P

Vosjan (1975), Tsuji and Yagi (1980)

Desulfovibrio gigas

P

Hatchikian et al. (1976), Hatchikian et al. (1978)

Desulfovibrio profundus

P

Bale et al. (1997)

Desulfovibrio senezii

P

Tsu et al. (1998)

Desulfovibrio cavernae

P

Sass and Cypionka (2004)

Desulfovibrio lacusfryxellense

P

Sattley and Madigan (2010)

Desulfobacter hydrogenophilus

P

Widdel (1987)

Desulfobulbus propionicus

P

Laanbroek et al. (1982)

Desulfonema limicola

P

Widdel et al. (1983)

Desulfuromonas acetoxidans

IA

Brugna et al. (1999)

Desulfurella multipotens

P

Miroshnichenko et al. (1994)

Desulfurella kamchatkensis

P

Miroshnichenko et al. (1998)

Geothermobacter ehrlichii

G

Strittmatter et al. (2009)

Geobacter sulfurreducens

P

Caccavo et al. (1994)

Geobacter hydrogenophilus

P

Coates et al. (2001)

Geobacter grbiciae

P

Coates et al. (2001)

Hippea maritima

P

Miroshnichenko et al. (1999)

Desulfacinum hydrothermale

P

Sievert and Kuever (2000)

Pelobacter acetylenicus

P

Stieb and Schink (1985)

Syntrophus aciditrophicus

P

Jackson et al. (1999)

Syntrophus buswellii

PA Mountfort et al. (1984), Wallrabenstein et al. (1994)

Syntrophobacter wolinii

P

Syntrophobacter pfennigii

PA Wallrabenstein et al. (1995)

Syntrophobacter fumaroxidans

I

de Bok et al. (2002)

Smithella propionica

P

Liu et al. (1999)

Boone and Bryant (1980), Wallrabenstein et al. (1994)

Helicobacter pylori

A

Maier et al. (1996a)

Nautilia lithotrophica

P

Miroshnichenko et al. (2002)

Wolinella succinogenes

P

Wolin et al. (1961)

I

Aspen and Wolin (1966), Dross et al. (1992)

Campylobacter jejuni

PA Laanbroek et al. (1982), Goodman and Hoffman (1983)

Campylobacter sputorum

P

Schumacher et al. (1992)

Sulfurospirillum deleyianum

P

Schumacher et al. (1992)

Sulfurospirillum arcachonense

P

Finster et al. (1997)

Sulfurospirillum paralvinellae

P

Takai et al. (2006)

Hydrogenimonas thermophila

P

Takai et al. (2004a)

Thermodesulfobacterium commune

P

Hatchikian and Zeikus (1983)

Thermodesufatator indicus

P

Moussard et al. (2004)

Geothermobacterium ferrireducens

P

Kashefi et al. (2002b)

Thermotoga maritima

P

Huber et al. (1986)

I

Juszczak et al. (1991), Vargas et al. (1998)

Thermotoga neapolitana

P

Jannasch et al. (1988)

Thermotoga thermarum

P

Windberger et al. (1989)

Thermotoga elfii

P

Ravot et al. (1995)

4

125

126

4

H2-Metabolizing Prokaryotes

. Table 4.1 (continued) Group

Incertae sedis

Order

Speciesa

Eb References

Sulfurimonas denitrificans

G

Sievert et al. (2008)

Oceanotoga teriensis

P

Jayasinghearachchi and Lal (2011)

Marinitoga camini

P

Wery et al. (2001)

Thermosipho africanus

P

Ravot et al. (1995)

Thermosipho atlanticus

P

Urios et al. (2004)

Thermosipho geolei

P

L’Haridon et al. (2001)

Fervidobacterium islandicum

P

Huber et al. (1990)

Caldithrix abyssi

P

Miroshnichenko et al. (2003a)

Abbreviations: E, evidence; I, hydrogenase has been isolated; P, physiological evidence for hydrogenase activity; G, genetic evidence; A, biochemical assay a Green: H2 producer; brown: H2 consumer; and blue: both b Type of evidence c Formerly Calderobacterium hydrogenophilum d Formerly Hydrogenobacter acidophilus e Formerly Anacystis nidulans (=SAG 1402–1) f =CCAP 1430/1, =CALU 743 g =CCAP 1403/13A, =ATCC 29413 h =CCAP 1403/2A, =SAG 1403–2, =ATCC 27899 i =CCAP 1427/1 j Formerly Clostridium thermoaceticum k Formerly Paracoccus denitrificans l Formerly Rhodocyclus gelatinosus m Formerly Desulfovibrio desulfuricans Norway 4 n Formerly Desulfovibrio baculatus

attracted much attention in recent years (Happe et al. 1994; Florin et al. 2001). Furthermore, some eukaryotic microbes possess organelles called ‘‘hydrogenosomes’’ which contain hydrogenases and engage in H2 metabolism (Mu¨ller 1993). Since organelles are descendants of prokaryotic cells, these organisms will be mentioned briefly but not treated in-depth. A number of review articles have been published on various aspects of H2 metabolism and hydrogenases (Adams 1990; Wu and Mandrand 1993; Sasikala et al. 1993; Albracht 1994; Vignais and Toussaint 1994; Fontecilla-Camps 1996; Maier and Bo¨ck 1996; Frey 1998; Nandi and Sengupta 1998; Peters 1999; Casalot and Rousset 2001; Vignais et al. 2001; Horner et al. 2002; Vignais and Billoud 2007). The reader is also referred to reviews on various groups of H2-metabolizing prokaryotes including sulfate reducers (Fauque et al. 1988; Voordouw 1995), aerobic H2-oxidizing bacteria (Aragno and Schlegel 1992; Friedrich and Schwartz 1993), methanogens (Sorgenfrei et al. 1997; Thauer 1998; Deppenmeier 2002; Thauer et al. 2010), homoacetogens (Drake 1994; Ragsdale and Pierce 2008), Sulfate reducers (Matias et al. 2005), and cyanobacteria (Tamagnini et al. 2002, 2007; Ghirardi et al. 2007).

Ecology The term ‘‘H2-metabolizing prokaryotes’’ lumps together many taxonomically and physiologically unrelated organisms. Accordingly, the representatives of this group are found in a range of very different habitats. This section does not attempt to describe exhaustively the biotopes of H2-metabolizing prokaryotes.

The emphasis, rather, is on the interactions between H2producers and H2-consumers, which are important for global geochemical cycles.

The Global H2 Budget Before considering the ecophysiology of the various groups of H2-metabolizing prokaryotes and the microbial communities they belong to, it is appropriate to briefly discuss the global budget of atmospheric H2. Three types of processes impact the pool of atmospheric H2: anthropogenic, biogenic, and geochemical. The total amount of H2 released annually into the troposphere has been estimated to be 107  15 Tg (Rhee et al. 2006). As will be discussed later, substantial amounts of H2 are turned over within microbial ecosystems without, however, affecting the atmospheric H2 budget. Conrad has summarized the numerous studies on atmospheric H2 (Conrad 1988). Anthropogenic activities are a major source of atmospheric H2. Equally important is the production of H2 by the oxidation of atmospheric methane (CH4) and nonmethane hydrocarbons. This entails chemical reactions with photochemically produced hydroxyl radical. Photochemical evolution of H2 accounts for 30–40 % of the total production. In contrast, the total biospheric emission, that is, the contribution of all ocean, lake, and soil biota to the global H2 pool, is only 7–11 %. Several studies have addressed the sources of biogenic H2 emission breaking it down into its marine, lacustrine, and terrestrial components. The major part of the euphotic surface layer of the world’s oceans is supersaturated for H2. Thus, a net emission of H2

H2-Metabolizing Prokaryotes

. Table 4.2 H2-metabolizing archaea Group

Order

Crenarchaeota Desulfurococcales

Thermoproteales

Sulfolobales

Euryarchaeota Archaeoglobales

Methanosarcinales

Speciesa

Eb References

Pyrodictium brockii

PA Stetter et al. (1983), Pihl et al. (1989) I

Pihl and Maier (1991)

Pyrodictium ocultum

P

Stetter et al. (1983), Fischer et al. (1983)

Pyrodictium abyssi

P

Pley et al. (1991)

Thermodiscus maritimus

P

Fischer et al. (1983)

Pyrolobus fumarii

P

Blo¨chl et al. (1997)

Hyperthermus butylicus

P

Zillig et al. (1990)

Ignicoccus islandicus

P

Huber et al. (2000a)

Ignicoccus pacificus

P

Huber et al. (2000a)

Ignicoccus hospitalis

P

Paper et al. (2007)

Stetteria hydrogenophila

P

Jochimsen et al. (1997)

Thermoproteus tenax

P

Zillig et al. (1981), Fischer et al. (1983)

Thermoproteus neutrophilus

P

Fischer et al. (1983)

Pyrobaculum islandicum

P

Huber et al. (1987)

Pyrobaculum aerophilum

P

Volkl et al. (1993)

Sulfolobus solfataricus

P

Brock et al. (1972)

Sulfolobus acidocaldarius

P

Brock et al. (1972)

Metallosphaera sedula

P

Huber et al. (1989)

Acidianus brierleyi

P

Segerer et al. (1986)

Acidianus infernus

P

Segerer et al. (1986)

Acidianus ambivalens

P

Stetter et al. (1986)

Stygiolobus azoricus

P

Segerer et al. (1991)

Archaeoglobus fulgidus

P

Stetter (1988), Klenk et al. (1997)

Archaeoglobus profundus

P

Burggraf et al. (1990b)

Archaeoglobus lithotrophicus

P

Stetter et al. (1993)

Archaeoglobus sulfaticallidus

P

Steinsbu et al. (2010)

Ferroglobus placidus

P

Hafenbradl et al. (1996)

Geoglobus acetivorans

P

Slobodkina et al. (2009a)

Methanosarcina barkeri Fusaro

I

Fiebig and Friedrich (1989)

G

Ku¨nkel et al. (1998), Vaupel and Thauer (1998), Meuer et al. 1999

P

Weimer and Zeikus (1978)

I

Fauque et al. (1984)

Methanosarcina barkeri DSM 800 Methanosarcina mazei Go¨1 Methanomicrobiales Methanospirillum hungatei

I

Mah (1980), Deppenmeier et al. (1992)

G

Deppenmeier et al. (1995), Deppenmeier (1995)

P

Ferry et al. (1974)

I

Sprott et al. (1987)

Methanocalculus halotolerans

P

Ollivier et al. (1998)

Methanomicrobium mobile

P

Paynter and Hungate (1968)

Methanocorpusculum parvum

P

Zellner et al. (1989)

Methanocorpusculum sinense

P

Zellner et al. (1989)

Methanocorpusculum bavaricum

P

Zellner et al. (1989)

Methanogenium organophilum

P

Widdel et al. (1988)

Methanogenium marisnigri

P

Romesser et al. (1979)

Methanogenium cariaci

P

Romesser et al. (1979)

Methanogenium frigidum

P

Franzmann et al. (1997)

4

127

128

4

H2-Metabolizing Prokaryotes

. Table 4.2 (continued) Group

Order

Methanobacteriales

Speciesa

Eb References

Methanoplanus limicola

P

Wildgruber et al. (1982)

Methanoplanus endosymbiosus

P

van Bruggen et al. (1986)

Methanothermobacter thermoautotrophicusc DH

P

Zeikus and Wolfe (1972), Jacobson et al. (1982)

I

Kojima et al. (1983), Fox et al. (1987)

G

Alex et al. (1990), Smith et al. (1997)

Methanothermobacter marburgensisd DSM 2133

Methanocellales Methanococcales

Methanopyrales Thermococcales

PA Zirngibl et al. (1990), Afting et al. (1998) I

Setzke et al. (1994)

G

Tersteegen and Hedderich (1999)

Methanobrevibacter smithii

P

Miller et al. (1986)

Methanobrevibacter acididurans

P

Savant et al. (2002)

Methanobacterium formicicum sp.

P

van Bruggen et al. (1984)

Methanobacterium formicicum MF

IA

Jin et al. (1983)

Methanobacterium formicicum JF-1

IA

Baron and Ferry (1989a), Baron and Ferry (1989b), Baron et al. (1989)

Methanobacterium alcaliphilum

P

Blotevogel et al. (1985), Worakit et al. (1986)

Methanobacterium aarhusense

P

Shlimon et al. (2004)

Methanothermus fervidus

P

Stetter et al. (1981)

Methanothermus sociabilis

P

Lauerer et al. (1986)

Methanosphaera stadtmanae

P

Miller and Wolin (1985)

Methanocella paludicola

P

Sakai et al. (2008)

Methanocaldococcus jannaschii

G

Sakai et al. (2011)

P

Jones et al. (1983a)

I

Shah and Clark (1990)

G

Halboth and Klein (1992), Bult et al. (1996)

Methanocaldococcus infernus

P

Jeanthon et al. (1998)

Methanocaldococcus vulcanius

P

Jeanthon et al. (1999)

Methanocaldococcus indicus

P

L’Haridon et al. (2003)

Methanothermococcus thermolithotrophicus

P

Huber et al. (1982), Belay et al. (1986)

Methanothermococcus okinawensis

P

Takai et al. (2002)

Methanococcus vanielli

I

Yamazaki (1982)

Methanococcus voltae

I

Muth et al. (1987)

Methanococcus maripaludis

P

Jones et al. (1983b)

G

Major et al. (2010)

Methanotorris igneus

P

Burggraf et al. (1990a)

Methanopyrus kandleri

P

Kurr et al. (1991)

G

Slesarev et al. (2002)

P

Fiala and Stetter (1986)

I

Bryant and Adams (1989), Ma et al. (1993), Sapra et al. (2000), Silva et al. 2000

Pyrococcus abyssi

P

Erauso et al. (1993)

Pyrococcus woesei

P

Zillig et al. (1987)

Palaeococcus ferrophilus

P

Takai et al. (2000)

Thermococcus litoralis

P

Neuner et al. (1990)

I

Ra´khely et al. (1999)

Pyrococcus furiosus

Thermococcus stetteri Thermococcus celer

P

Miroshnichenko et al. (1989), Pusheva et al. (1991)

I

Zorin et al. (1996a)

P

Zillig et al. (1983)

H2-Metabolizing Prokaryotes

4

. Table 4.2 (continued) Group

Order

Speciesa

Eb References

Thermococcus thioreducens

P

Pikuta et al. (2007)

Thermococcus kodakaraensis

P

Atomi et al. (2004)

Abbreviations: E, evidence; I, hydrogenase has been isolated; P, physiological evidence for hydrogenase activity; G, genetic evidence; A, biochemical assay Green: H2 producer; brown: H2 consumer; and blue: both b Type of evidence c Formerly Methanobacterium thermoautotrophicum DH d Formerly Methanobacterium thermoautotrophicum Marburg a

from the surface water to the atmosphere must take place. While experimental data are still lacking, the main source of this H2 is probably the N2-fixing cyanobacteria and prochlorophytes. The same probably holds true for H2 production in the oxic layers of freshwater lakes. The H2 concentration of the epilimnion reaches levels of 0.5–50 nM (Schropp et al. 1987). Studies have shown that the H2 concentration of lake water correlates with cell counts of cyanobacteria on the one hand and with N2 fixation rates on the other (Conrad et al. 1983a; Schmidt and Conrad 1993; Schu¨tz et al. 1988). The results refute the older notion that H2 arises by fermentation in the anoxic sediment and diffuses up into the oxic zone. Almost all of the H2 evolved in anoxic sediments is also consumed there. The contribution of soil to the atmospheric H2 pool is more complex. Most soils do not emit H2 but, on the contrary, consume it (Seiler 1978; Conrad and Seiler 1981, 1985). An exception to this are soils in areas where leguminous plants grow. Symbiotic rhizobia in root nodules produce H2 in conjunction with N2 fixation. An estimated 1 million tons of H2 is produced by nodule bacteria annually (Evans et al. 1987). During the vegetation period, the rates of H2 production by root nodules are high enough to lead to a net increase in the H2 concentration in the soil. A portion of this H2 escapes to the atmosphere. Thus, in all three major environmental zones, release of H2 into the atmosphere is a result of N2 fixation. The production of H2 by microbial fermentation processes in the gut of termites may be an exception to this generalization. It has been estimated that as much as 1014 g of H2 could be released into the atmosphere annually by these microbes (Zimmerman et al. 1982). Experimental data on this question are inconclusive. Laboratory experiments with termites pointed to a significant release (Zimmerman et al. 1982). On the other hand, measurements made in the field on actual termite mounds showed that there was no measurable release of H2 at all (cited by Conrad (1988) as ‘‘W. Seiler and R. Conrad, unpublished observation’’). On the consumption side of the balance sheet, chemical processes in the atmosphere are responsible for only a small fraction of H2 decomposition. By far, the most important global sink for H2 is the soil, which accounts for over 90 % of the total global consumption (Rhee et al. 2006). Remarkably, consumption of atmospheric H2 in soils may be to a significant extent only indirectly attributable to microorganisms. Rather, it appears that H2-oxidizing activity associated with soil particles is at least in part the basis of H2 consumption by soil (reviewed by Conrad 1996). Various lines of evidence support this conclusion:

(1) For an organism to utilize atmospheric H2, it must have a Km for H2 in the range of 5–80 nM (Conrad 1984). Several characterized strains of chemolithotrophic bacteria have Km values for H2 above 0.5 mM. (2) Suspensions of H2-oxidizing chemolithotrophs provided with a mixture of H2 and air as growth substrate consume H2 down to a certain concentration. After this point, no more H2 is utilized and growth ceases. The critical concentration for H2 uptake is known as the threshold value. The threshold for H2 utilization for various H2oxidizing laboratory strains (1–10 p.p.m.v.) is significantly higher than the concentration of H2 in the atmosphere (0.5 p.p.m.v.) (Conrad et al. 1983b; Conrad and Seiler 1979; Schuler and Conrad 1990). (3) The H2-oxidizing activity of the soil is destroyed by boiling and autoclaving and has an optimum of 25–40  C (Seiler 1978; Fallon 1982; Schuler and Conrad 1991a). (4) The H2-oxidizing activity associated with sizefractioned soil particles is not correlated with parameters indicative of microbial biomass (ATP content and microscopic cell counts; Ha¨ring et al. 1994). (5) Treatment of soil samples with chloroform or sodium azide caused a loss of H2 uptake activity but did not abolish it altogether. Taken together, these results suggest that organisms are present in the soil, which are capable of oxidizing atmospheric H2. The putative high-affinity hydrogenase enzymes released from lysed bacteria and immobilized on soil particles or persisting in dead cells could also contribute to the oxidation of atmospheric H2. Indeed, an actinobacterial strain designated Streptomyces sp. PCB7 was recently isolated from a soil sample. This strain is capable of utilizing H2 down to a threshold concentration of 0.1 p.p.m.v. (Constant et al. 2008). This strain carries genes for a novel type of [NiFe] hydrogenase. A PCR-based survey of samples from a wide variety of soils indicated that organisms equipped with this type of hydrogenase are ubiquitous (Constant et al. 2011). Many ‘‘classical’’ H2-oxidizing chemolithoautotrophs are readily isolated from a variety of soil biotopes. Why are chemolithoautotrophs widespread in soils if they are not able to metabolize atmospheric H2? The following aspects are important in this context and may be at least part of the explanation: (1) Most of the aerobic, mesophilic H2-oxidizers (knallgas bacteria) isolated from soils are facultative chemolithoautotrophs that also thrive organotrophically (Aragno and Schlegel 1992). Such organisms are predestined to utilize H2 that is transiently available in biologically relevant concentrations. It is well known that soils rapidly become anoxic when they are waterlogged. This can lead to a transient production of H2 when soil microbes shift

129

130

4

H2-Metabolizing Prokaryotes

to fermentation. (2) Various soil microenvironments, for example, the vicinity of root nodules, may provide high local concentrations of H2 which could at least transiently support the growth of H2-oxidizing chemolithotrophs. While many symbiotic N2-fixing bacteria have uptake hydrogenase and hence can oxidize the H2 produced by their nitrogenases, over 50 % of diazotrophs lack hydrogenases (Arp 1992). The so-called Hupnegative (Hup ) strains are responsible for the substantial amounts of H2 released by root nodules. Legumes nodulated with Hup -rhizobia may release as much as 5,000 L per day and hectare (Dong et al. 2003). Most of this H2 is oxidized in the immediate vicinity of the nodule, and very little of it escapes into the atmosphere (Conrad and Seiler 1979; La Favre and Focht 1983; Cunningham et al. 1986). A recent comparative study surveyed the population of H2-oxidizing chemolithotrophs associated with Hup+- and Hup -nodulated legumes (Maimaiti et al. 2007). In soil samples taken from fields with Hup -nodulated plants, about 1 % of the aerobic isolates could oxidize H2 at concentrations typical for chemolithotrophs. No H2-oxidizing strains were obtained from fields of Hup+-nodulated plants. (3) Syntrophic associations between H2 producers and H2 consumers may be a widespread phenomenon. Juxtaposition of individual cells would, in effect, provide the consumer with a high local concentration of H2. This will be discussed in more detail below.

Habitats of H2-Evolving Prokaryotes Anaerobic food chains which degrade organic material via the various fermentation processes outlined above are a major source of H2 in the biosphere. Fermenting organisms are limited to anoxic zones rich in organic substance. Marine and lacustrine sediments are the most important biotopes of this sort. These sediments are fed by a constant influx of organic material derived from photosynthetic primary producers and from the ensuing food chains. The upper, oxic layer of the sediment varies in depth both in marine and freshwater sediments. Below this layer is the zone of anoxic decomposition. In this stratum, H2 is evolved as a product of fermentation. The H2 produced neither accumulates nor does it escape in significant quantities to the oxic zone. If H2 were to accumulate, the fermentative metabolic processes would soon come to a halt, since these are inhibited by relatively low concentrations of H2 in the environment. The inhibitory concentrations vary for the different fermentative reactions, depending on their energetics. Fermentation of fatty acids to acetate, H2, and CO2, for instance, is more endergonic than the fermentation of ethanol to acetate and H2, and the former process ceases at a much lower concentration of external H2 than the latter (see Chap. 21, ‘‘Syntrophism Among Prokaryotes’’ in Vol. 2). Studies on the sediment of a eutrophic lake showed that fermentation of butyrate and propionate was inhibited by H2 concentrations of 100 and 20 nM, respectively (Conrad et al. 1986). The concentration of H2 is kept at a low level (i.e., 5–30 nM in marine sediments and 1–150 nM in lacustrine sediments) due to constant depletion by

H2-consuming organisms (reviewed by Jorgensen 1989; see also Strayer and Tiedje 1978; Lovley and Klug 1982; Robinson and Tiedje 1982; Phelps and Zeikus 1984; Conrad et al. 1985). High turnover rates for H2 have been measured. In a study on a lake sediment, the turnover time of the pool of free H2 was estimated to be 2 min (Conrad et al. 1985). Two minor habitats of fermenting bacteria deserve mention: the stomach of ruminants and the gut of xylophagous arthropods including termites (Breznak 1982; Zimmerman et al. 1982; Leschine 1995; Ricke et al. 1996; Flint 1997). Spirochetes have been isolated from the termite hindgut. While some of these isolates are homoacetogens that consume H2, others ferment carbohydrates and produce H2 (Graber et al. 2004). The rumen harbors a rich microbial flora. Among these microorganisms are bacteria such as Ruminococcus albus and Butyrivibrio fibrisolvens which ferment cellulose to organic acids, H2, and CO2. Hydrogen is generated as a byproduct of N2 fixation in both oxic and anoxic environments. Cyanobacteria and prochlorophytes are probably the most widespread diazotrophs on earth. These organisms inhabit the upper, oxic zones of oceans and lakes. The anoxygenic photosynthetic bacteria, which occupy deeper zones depleted for O2, also engage in N2 fixation. These and other diazotrophs contain uptake hydrogenases and, hence, are capable of exploiting at least a part of the H2 generated by nitrogenase. However, hydrogenase-free strains abound in nature resulting in the liberation of large quantities of H2. Endosymbiotic rhizobia are the third important group of diazotrophs. The role of these organisms in rhizospheric H2 production has been mentioned above. The extensive mats found in coastal areas are a special cyanobacterial habitat. The mats consist of a gelatinous mass produced by the microbes, and the generation of H2 leads to the formation of bubbles in this viscous matrix. The mats formed by different microbial communities differ in their consistency. This, in turn, determines whether the H2 is retained or released into the atmosphere (Hoehler et al. 2001).

Habitats of H2-Consuming Prokaryotes Both biogenic and abiogenic H2 production can support growth of H2-utilizing prokaryotes. Many H2-utilizing species profit from fermentative H2 production in anoxic sediments. The two most important groups of H2-consumers in such biotopes are the methanogenic archaea and the sulfate-reducing bacteria. It has been known for some time that methanogens and sulfate reducers compete for H2 (Winfrey and Zeikus 1977; Winfrey et al. 1977; Abram and Nedwell 1978; Oremland and Polcin 1982; Lovley et al. 1982; Lovley and Klug 1983). Sulfate reducers outcompete the methanogens in the presence of sulfate because the former have a higher affinity for H2 and a higher growth yield (Kristjansson et al. 1982; Scho¨nheit et al. 1982). In studies using pure cultures, Km and YH2 values of 5.0  0.5 mM and 0.2 g protein/mol H2, respectively, were determined for Methanospirillum hungatei JF-1 versus 1.1  0.1 mM and 0.85 g protein/mol H2, respectively, for Desulfovibrio strain G11

H2-Metabolizing Prokaryotes

(Robinson and Tiedje 1984). The key factor determining which of the two terminal degradation processes—sulfidogenesis or methanogenesis—prevails in a given habitat is SO42 concentration. In anoxic marine sediments, where there is an abundant supply of SO42 , sulfate reduction is the dominant process, consuming most of the available H2 and acetate. The sulfate level in lakes varies depending on their trophic state but in general is lower than in seawater. The thickness of the zone of sulfate reduction varies accordingly. In eutrophic lakes, the sulfate concentration in the sediment drops of sharply. Here, the zone of sulfate reduction is only a few centimeters thick. In lakes with lower nutrient contents, the zone of sulfate reduction extends deeper into the sediment (Lovley and Klug 1983). Competition for H2 does not mean that sulfate reduction and methanogenesis are mutually exclusive processes. Various methanogens can exploit substrates, for example, methylamine, that are not utilized by sulfate reducers (Oremland and Polcin 1982; Winfrey and Ward 1983). Therefore, the two groups of organisms can coexist in the same biotope, as has been shown for instance for estuarine sediments. According to one estimate based on sediment from an oligotrophic lake, the fraction of the total flux of electrons and carbon routed through sulfate reduction is between 30 % and 81 % of the total terminal metabolism (Lovley and Klug 1983). Sulfate reduction is also an important process in extreme environments, such as the anaerobic sediments of soda lakes. Among the specialized, H2-utilizing, sulfate-reducing bacteria found in such sediments are the alkaliphilic lithoheterotroph Desulfonatronovibrio hydrogenovorans and the alkaliphilic lithoautotrophs Desulfonatronum lacustre (Zhilina et al. 1997; Pikuta et al. 1998) and Desulfonatronum cooperativum (Zhilina et al. 2005). The role of soils as habitats for facultative H2-oxidizing chemolithotrophs has been discussed above. Two niches are especially important in this context: the rhizosphere in the vicinity of nodulated plants and the interface of anoxic enclaves where H2 is evolved and can diffuse into the surrounding oxic zone. Several obligately chemolithoautotrophic bacteria, which oxidize either H2 or sulfur, have been described. These include both mesophilic and thermophilic forms. The latter are mostly confined to special niches such as hot springs and hydrothermal vents (see below for a detailed discussion of hydrothermal vents). Hydrogenobacter thermophilus and Hydrogenobacter halophilus inhabit freshwater and saline hot springs, respectively (Kawasumi et al. 1984; Nishihara et al. 1989, 1990). The obligate H2-oxidizing Fe(III) reducers Geothermobacterium ferrireducens (Kashefi et al. 2002) and Thermolithobacter ferrireducens (Sokolova et al. 2007) were isolated from the sediments of terrestrial hot springs. The hyperthermophilic H2-oxidizing bacterium Aquifex pyrophilus grows in hot marine sediments (Huber et al. 1992). Several obligately chemolithotrophic archaea have been identified. Thermodiscus maritimus and Pyrodictium occultum inhabit submarine solfataric springs (Fischer et al. 1983). Two obligate H2-oxidizing, sulfatereducing archaea, Ignicoccus islandicus and Ignicoccus pacificus,

4

were enriched from hot marine sediments and from the orifice of a deep-sea vent, respectively (Huber et al. 2000b). These organisms utilize the H2, CO2, and sulfate dissolved in the hydrothermal fluid for growth. Thus, they are chemolithoautotrophic primary producers and form the basis of food chains in their respective habitats (Jannasch and Mottl 1985). The hyperthermophile Archaeoglobus profundus is another example of obligate H2-based lithotrophy (Burggraf et al. 1990b). However, this organism requires organic carbon sources.

Syntrophy and Interspecies H2 Transfer Syntrophy is the mutual metabolic dependence of two different types of prokaryotes (see Chap. 21, ‘‘Syntrophism Among Prokaryotes’’ in Vol. 2). The first obligately syntrophic relationship involving an exchange of H2 between the partner organisms was recognized by Bryant (Bryant et al. 1967). He discovered that the ethanol-degrading ‘‘bacterium’’ Methanobacillus omelianskii was in fact a coculture of a so-called S-organism, which was the actual ethanol-degrader, and a methanogen. The ethanoldegrader was strictly dependent on the methanogen, because the latter consumed H2, thereby ‘‘pulling’’ the otherwise thermodynamically unfavorable oxidation of ethanol. Wolin reported new examples of microbial H2 exchange and introduced the phrase ‘‘interspecies hydrogen transfer’’ to describe this general phenomenon (Scheifinger et al. 1975; Wolin 1976, 1982). Based on quantitative studies on freshwater sediments, Conrad proposed the juxtaposition of H2-producers and H2-consumers (Conrad et al. 1985). He suggested that the two types of cells are in close, physical contact in particles or flocs. As a result of juxtaposition, a major fraction of the H2 produced would never enter the pool of dissolved H2 but rather be transferred directly from cell to cell. This would explain the discrepancy between turnover rates in the extracellular H2 pool and growth yield of the population of methanogens. The interspecies transfer of H2 between juxtaposed cells of H2-producers and H2-consumers is especially important for organisms which ferment fatty acids, such as butyrate and propionate, since for thermodynamic reasons these processes are inhibited by low levels of H2 (Boone and Bryant 1980). Numerous syntrophic H2-producing strains have been identified as partners in interspecies H2 transfer. These include strains such as Desulfovibrio vulgaris, Thermoanaerobacter brockii, and Pelobacter venetianus which ferment primary alcohols (Bryant et al. 1977; Ben-Bassat et al. 1981; Schink and Stieb 1983), the butyrate and propionate fermenters Syntrophomonas wolfei, Syntrophomonas bryantii, Syntrophobacter wolinii, Syntrophobacter pfennigii, and Smithella propionica (McInerney et al. 1979; Boone and Bryant 1980; Stieb and Schink 1985; Wallrabenstein et al. 1995; Liu et al. 1999), the acetate oxidizer Thermacetogenium phaeum (Hattori et al. 2000), and the oligosaccharide-fermenting strain Thermicanus aegypticus (Go¨ssner et al. 1999). Interestingly, the propionate fermenters isolated so far are all capable of reducing sulfate (Harmsen et al. 1995; Conrad 1999). On the other hand, there is important indirect

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H2-Metabolizing Prokaryotes

evidence that a significant portion of the syntrophic population of some sediments is made up of sulfate reducers. Addition of sulfate to freshwater methanogenic sediments caused an immediate cessation of methane production (Lovley et al. 1982; Conrad et al. 1987). The most likely explanation for this observation is that sulfate reducers, which at low-sulfate concentrations obtain energy mainly via fermentation and therefore release H2 to rid themselves of excess reductant, immediately switch to respiration when sulfate is available as the terminal electron acceptor. In doing so, they cut off the supply of H2 to the methanogens. A similar effect has been observed in methanogenic soils of rice paddies (Achtnich et al. 1995a, b; Krylova et al. 1997). Sulfate reducers are probably predestined to participate in syntrophic associations because of the ambivalent nature of their H2 metabolism: They can take on the role of H2-consumer or H2-producer, depending on the sulfate concentration in the environment (Boone and Bryant 1980; McInerney et al. 1981b; Traore et al. 1981).

H2-Based Ecosystems Aquifers Since intraterrestial habitats first came under microbiological scrutiny, numerous studies have documented microbial life at depths of thousands of meters within the Earth’s crust. These investigations have led to the realization that the deep subsurface constitutes a third realm of the biosphere alongside oceans and land surfaces. Although population densities of intraterrestrial microorganisms are low, they represent on aggregate a huge amount of biomass. According to one estimate, the total amount of carbon in subsurface microbiota may be as large as the carbon content of terrestrial and marine plants combined (Whitman et al. 1998). Investigations on samples sterilely extracted from solid drilling cores from basement rock at depths >1 km revealed that viable microbes exist in the mm-scale pores of both granitic and basaltic rock (Lehman et al. 2001; Gihring et al. 2006; Sahl et al. 2008). More important in terms of biomass and geochemical cycling are the microbiota of deep groundwater. Aquifers, the water-conducting strata of sedimentary rock and the water-filled fracture zones of igneous rock, represent more or less discrete habitats. Aquifers host different microbial ecosystems depending on their physical and chemical characteristics including temperature, pH, and concentrations of solutes. Aquifers are recharged from surface water and, hence, receive a certain input of dissolved organic carbon (DOC). This results in zones of heterotrophic metabolism, with the decomposition of organic matter following a sequence of terminal electron acceptors similar to the situation in soil sediments (Lovley and Chapelle 1996). In contrast, other zones are disconnected from the surface and its photosynthetic production of biomass and, thus, have the potential to host lithoautotrophic microbial communities. Sampling aquifer microbiota, which consist of both planktonic and attached forms, is technically challenging, and contamination by surface microbes poses a major problem

(Pedersen 1997). In recent years, careful surveys based on libraries of 16S rRNA sequences have provided inventories of the microorganisms living in deep aquifers. Studies of this kind have provided evidence for the existence of subsurface microbial ecosystems that depend on H2-based lithoautotrophic primary production. Such habitats are characterized by anoxic conditions, low DOC values, and a supply of geologically produced H2 and CO2. H2 arises in the Earth’s crust via various processes including volcanic activity, radiolysis of water, the decomposition of methane at high temperatures, and serpentinization in mafic rocks. One candidate for an H2-based ecosystem is the Lidy Hot Springs Aquifer in the Northwestern United States (Chapelle et al. 2002). The water, sampled at a depth of 200 m, had a temperature of 58  C and a low DOC content ( Fig. 4.1) characterized by different physicochemical conditions: the chimney-like concretions formed by the precipitation

4

of minerals dissolved in the hydrothermal fluid, the plume or cloud where the superheated fluid mixes with cold seawater, sediments in the vicinity of the chimneys, and colonies of invertebrates such as the tube-forming invertebrate worms. The chimneys themselves host numerous microhabitats. Between the outer and inner walls of the chimney steep gradients of temperature (0–350  C), O2 concentration and pH prevail. The porous structure of the chimney wall probably provides the structural basis for a continuum of varied niches for microbial life. In addition, the visible surfaces of the chimney are often occupied by dense mats of microorganisms. The microbial communities at the different vent fields are far from uniform. Rather the distribution of organisms is largely dependent on the varying composition of the hydrothermal fluid. This, in turn, is determined by the local geology of the underlying crust. In general, H2-rich (>10 mM) exudates are associated with vents located in ultramafic rock formations, whereas basaltic formations yield H2-poor ( Fig. 4.1). In light of the results of the cultureindependent surveys, it is not surprising that several hydrogenotrophic methanogens were discovered, such as Methanothermococcus okinawensis (Takai et al. 2002), Methanocaldococcus indicus (L’Haridon et al. 2003), and Methanotorris formicicus (Takai et al. 2004b). Novel representatives of the Archaeoglobales, for example, Geoglobus ahangari (Kashefi et al. 2002a), coupled H2 oxidation with Fe(III) reduction. A surprising finding of molecular phylogenetic surveys was the abundance of Epsilonproteobacteria in a wide variety of marine hydrothermal settings (Moyer et al. 1995; Reysenbach et al. 2000; Corre et al. 2001; Longnecker and Reysenbach 2001; Campbell et al. 2006). Numerous representatives of these versatile bacteria have now been cultivated and described. These include obligate and facultative H2-oxidizers such as Nautilia lithotrophica (Miroshnichenko et al. 2002), Caminibacter profundus (Miroshnichenko et al. 2004), Caminibacter mediatlanticus (Voordeckers et al. 2005), Hydrogenimonas thermophilus (Takai et al. 2004a), Lebetimonas acidiphila (Takai et al. 2005), Nitratifractor salsuginis, Nitratiruptor tergarcus (Nakagawa et al. 2005b), and Thioreductor micantisoli (Nakagawa et al. 2005a). The flexible use of different electron

4

donors and acceptors allows vent Epsilonproteobacteria to adapt to variable conditions in the mixing zone in and around vent chimneys. Sulfurovum sp. NBC37-1, for instance, can utilize H2, S0, and S2O32 as electron donors (Yamamoto et al. 2010). Nautilia nitratireducens oxidizes H2 with NO3 , S0, or SeO42 as terminal acceptor (Perez-Rodriguez et al. 2010). Several novel Aquificales were also isolated, such as Balnearium lithotrophicum (Takai et al. 2003b), Persephonella hydrogeniphila (Nakagawa et al. 2003), Persephonella marina (Go¨tz et al. 2002), and Thermovibrio ruber (Huber et al. 2002).

Physiology: Varieties of H2 Metabolism This section is a brief summary of the major metabolic activities involving H2 in prokaryotes. In light of the diversity of the metabolic types, it is obvious that this discussion is intended as an overview emphasizing the various physiological roles assumed by hydrogenases.

H2-Evolving Processes Fermentation The decomposition of organic matter via fermentation is one of the major biotic energy-yielding processes in anaerobic habitats (reviewed by > Chap. 13, ‘‘The Anaerobic Way of Life’’ in Vol. 2). Various types of fermentation result in the formation of H2 as a terminal product and hence constitute a substantial contribution to the global H2 balance. Both obligate and facultative fermenters produce H2. One of the best-studied representatives of the former group is Clostridium pasteurianum. This bacterium ferments glucose and other substrates and evolves H2 as a means of dispersing excess reductant. Hydrogen production is catalyzed by two monomeric [FeFe] hydrogenases (Chen and Mortenson 1974; Adams et al. 1989; Adams 1990). A classical example of fermentative H2 production in a facultative fermenter is mixed-acid fermentation in E. coli. This organism produces H2 via the formate hydrogenlyase reaction (reviewed by Sawers 1994). The so-called hydrogenase-3, one of four [NiFe] hydrogenase isoenzymes in E. coli, is part of a membrane-bound complex containing the enzyme formate dehydrogenase (Sawers et al. 1985; Sawers and Boxer 1986; Bo¨hm et al. 1990; Sauter et al. 1992). The formate hydrogenlyase complex converts formate, an intermediary fermentation product, to the gaseous products CO2 and H2 (> Fig. 4.2). Fermentative bacteria constitute a major group of rumen flora, and as such, they are instrumental in the breakdown of cellulose and other biopolymers (reviewed by Hungate 1966 and Flint 1997). Representatives of this group such as Butyrivibrio fibrisolvens, Ruminococcus albus, Megasphaera elsdenii, and Eubacterium limosum ferment various substrates to organic acids, CO2 and H2 (Miller and Wolin 1973, 1979; Joyner et al. 1977). Hydrogen does not accumulate in the digestive tract, however, since it is immediately consumed by methanogens

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H2-Metabolizing Prokaryotes

Periplasmic hydrogenase (D. gigas) H2 2H+

Membrane-bound hydrogenase (R. eutropha) H2

[NiFe] HydB

Hydrogenase-2 (E. coli ) H2

[NiFe]

2H+

Fe-S HydA

H2

[NiFe] HybC

HoxG

Cyt c

Membrane-accociated hydrogenase (T. roseopersicina)

2H+

Fe-S HoxK

HynL

Fe-S Fe-S Hyba HybO

Cyt b HoxZ

[NiFe]

2H+ QH2

HybB

Q

Fe-S HynS Cyt b Isp1

FdrC

Isp2

FdrB

Formate hydrogenlyase-1 (E. coli)

?

ΔµH+ HycC Fe-S HycG HycD Fe-S HycF Fe-S HycB

H2

[NiFe]

2H+

HycE

2H+

MoSe FdhF

H+

FAD FdrA

Bifurcating hydrogenase (T. maritima)

H2

+ CO2

[FeFe] Fe-S HydA

Fe-S FMN HydB

fumarate + 2H+

NAD+

Fe-S HydC

Soluble hydrogenase (R. eutropha)

formate Fdox

succinate

NADH

H2

Fdred

2H+

[NiFe] HoxH

Fe-S HoxU Fe-S FMN HoxY

CO-oxidizing:H2-forming complex (C. hydrogenoformans)

? ΔµNa+ /ΔµH+

H2

CooK Fe-S CooL

Out

In

[NiFe] CooH Fe-S CooX

Fe-S CooF Fe-S CooF

NAD+ NADH

HoxI HoxI

Soluble hydrogenase-1 (P. furiosus)

2H+ CooM CooU

Fe-S FMN HoxF

2H+

NiFeS Fe-S CooS

2H+ + CO2

NiFeS Fe-S CooS

CO + H2O

[NiFe] HydA

H2

Fe-S HydD Fe-S HydB

Fe-S FAD HydG

NADPH NADP+

. Fig. 4.2 Representatives of major classes of bacterial hydrogenases: subunit composition, catalytic activity, and cellular location. See text for details

and/or homoacetogens (Sharak-Genthner and Bryant 1987). Megasphaera elsdenii is with respect to H2 metabolism the best-studied organism of this group. It produces a monomeric [FeFe] hydrogenase which is one of the smallest known representatives (Van Dijk et al. 1979; Filipiak et al. 1989; Atta and Meyer 2000). The thermophilic archaea Pyrococcus furiosus and Thermococcus litoralis obtain energy via a strictly anaerobic metabolism and dispose of surplus reducing equivalents in the form of H2 or H2S (Malik et al. 1989; Bryant and Adams 1989; Blumentals et al. 1990; Scha¨fer and Scho¨nheit 1991; Ma et al. 1993; Ma and Adams 1994; Pedroni et al. 1995; Ra´khely et al. 1999). P. furiosus synthesizes two cytoplasmic hydrogenase isoenzymes (SHI and SHII) encoded by duplicated genes (Bryant and Adams 1989; Ma et al. 1993, 2000) and a third hydrogenase (MBH) that is clearly related to the multisubunit [NiFe] hydrogenase-3 of E. coli (Sapra et al. 2000). Studies on individual knockout mutants for the three enzymes and various combinations of them have helped to clarify their physiological roles. In the absence of S0, the MBH is the only enzyme capable of evolving

H2 and thus regenerating the pool of oxidized electron carriers to support growth (Schut et al. 2012). A complex isolated from P. furiosus membranes under mild conditions was shown to possess H2-evolving activity using reduced P. furiosus ferredoxin as electron donor (Silva et al. 2000). There is evidence indicating that the complex engages in proton pumping. This means that the reduction of protons by the P. furiosus MBH constitutes a simple form of anaerobic respiration (Jenney and Adams 2008). The role of the cytoplasmic enzymes is still not clear. The thermophile Thermococcus litoralis seems to have a similar complement of hydrogenases. A tetrameric, cytoplasmic enzyme has been characterized (Ra´khely et al. 1999). A membrane-bound, ECH-type hydrogenase is part of formate hydrogenlyase complex similar to that found in E. coli (Taka´cs et al. 2008). Another example of fermentative H2 production is found in Thermotoga maritima, a thermophilic, strictly anaerobic bacterium living in warm marine sediments (Huber et al. 1986; Conners et al. 2006). Fermentative H2 evolution in T. maritima is catalyzed by a heterotrimeric, cytoplasmic [FeFe] hydrogenase (Verhagen et al. 1999; Jenney and Adams 2008). For many

H2-Metabolizing Prokaryotes

years, the physiological redox carrier of T. maritima hydrogenase was not known. Neither reduced T. maritima ferredoxin nor NADPH functioned as electron donors in vitro (Blamey et al. 1994; Verhagen et al. 1999). Schut and Adams solved this enigma by showing that both electron carriers are required simultaneously. The enzyme oxidizes a ferredoxin and an NADPH concomitantly, channeling the electrons to H2 (Schut and Adams 2009). The striking feature of this reaction is the coupling of an energetically unfavorable redox reaction with an energetically favorable one. To emphasize this characteristic feature, Schut and Adams proposed a novel class of hydrogenases designated ‘‘bifurcating [FeFe] hydrogenases’’ (> Fig. 4.2). An important group of specialized fermenters are the syntrophic, fatty-acid oxidizers which inhabit anaerobic sediments (reviewed by Chap. 21, ‘‘Syntrophism among Prokaryotes’’ in Vol. 2). These bacteria ferment butyrate and propionate via reactions that are endergonic under standard conditions (Thauer et al. 1977): CH3 CH2 CH2 COOH þ 2H2 O ! 2CH3 COOH þ 2H2 ð4:3Þ CH3 CH2 COOH þ 2H2 O ! 2CH3 COOH þ CO2 þ 3H2

ð4:4Þ

To solve the thermodynamic problem inherent in these reactions, the syntrophic fermenters live in aggregates with H2-utilizing organisms such as methanogens. Hydrogen is consumed as rapidly as it is produced, preventing its accumulation in the milieu. Thus, the reactions are ‘‘pulled’’ in the direction of the oxidized products. The cyanobacteria are strictly phototrophic organisms. However, many cyanobacterial strains switch to a fermentative metabolism during periods of darkness when O2 of the milieu is exhausted by respiration (reviewed in Stal and Moezelaar 1997). In this metabolic state, they usually consume endogenous reserves (such as glycogen) that accrue during photosynthetic growth. In Microcystis PCC7806 and Cyanothece PCC7822, for instance, endogenous glycogen is fermented to ethanol, acetate, lactate, CO2, and H2 via a mixed-acid pathway using ferredoxin as oxidant (van der Oost et al. 1989; Moezelaar and Stal 1994). Hydrogenase couples H2 production to ferredoxin reoxidation. Microcoleus chthonoplastes employs a similar mixedacid-type fermentation to generate energy under anoxic conditions (Moezelaar et al. 1996). However, this organism produces hydrogen from formate via a formate hydrogenlyase reaction. Many anoxygenic phototrophs ferment endogenous reserves in the dark and thereby produce H2 as one of the fermentation products. Rhodospirillum rubrum grows anaerobically in the dark on fructose or pyruvate (Scho¨n 1968; Uffen and Wolfe 1970; Scho¨n and Biedermann 1973; Uffen 1973a, b; Jungermann and Scho¨n 1974). The key enzyme for the fermentation of pyruvate is pyruvate-formate lyase (Gorrell and Uffen 1977, 1978). Formate hydrogenlyase catalyzes the evolution of CO2 and H2 from formate (Kohlmiller and Gest 1951). Rhodobacter capsulatus ferments fructose to succinate, lactate, acetate, CO2, and H2 (Yen and Marrs 1977; Madigan and Gest 1978; Schultz and Weaver 1982). Also capable of fermentation are

4

Rhodopseudomonas palustris, Rhodopseudomonas viridis, and Rhodobacter sphaeroides (Uffen and Wolfe 1970). Thiocapsa roseopersicina evolves H2 during fermentative growth in the dark and in the presence of glucose at low sodium thiosulfate levels. Two cytoplasmic [NiFe] hydrogenases are responsible for H2 evolution (Ra´khely et al. 2007; Maro´ti et al. 2010).

Anaerobic CO Oxidation The anoxygenic photosynthetic bacterium Rhodospirillum rubrum can grow anaerobically in the dark on CO as the sole source of energy (Uffen 1976). Under these conditions, it forms a membrane-bound enzyme complex consisting of carbon monoxide dehydrogenase (CODH) and a CO-insensitive [NiFe] hydrogenase (Fox et al. 1996a, b). Together, these enzymes catalyze the following net reaction: CO þ H2 O ! CO2 þ H2

ð4:5Þ

Like the hydrogenase-3 of E. coli, this multienzyme complex includes other proteins related to reduced nicotinamide adenine dinucleotide (NADH):quinone dehydrogenase (complex I; Fox et al. 1996a). The fact that CO supports growth of R. rubrum in the dark argues for an energy-conserving function of the CO dehydrogenase-hydrogenase complex. Those carboxydotrophs which couple the oxidation of CO to the reduction of protons are known as hydrogenogens (Svetlitchnyi et al. 2001; Oelgeschla¨ger and Rother 2008; Sokolova et al. 2009). A well-studied representative of this group is the strictly anaerobic, thermophilic, Gram-positive bacterium Carboxydothermus hydrogenoformans (Svetlichnyi et al. 1991; Soboh et al. 2002). Genetic determinants for five different CODHs have been identified in the genome sequence of this organism (Wu et al. 2005). One of them, designated CODH I, forms a multienzyme membrane-bound complex with a hexameric Ech-type hydrogenase (> Fig. 4.2; Svetlitchnyi et al. 2001; Soboh et al. 2002). The energetic basis for growth on CO is the proton-pumping activity of the CODH-ECH complex in conjunction with chemiosmotic ATP synthesis. Included in the ranks of the thermophilic hydrogenogens are Carboxydocella thermautotrophica (Sokolova et al. 2002), Thermosinus carboxydivorans (Sokolova et al. 2004), Thermincola carboxydiphila (Sokolova et al. 2005), and Thermolithobacter carboxydivorans (Sokolova et al. 2007).

Production of H2 as a Byproduct of N2 Fixation Nitrogen fixation is one of the main processes of biogenic H2 production but is unique for the reason that it does not involve a specialized H2-forming enzyme, that is, a hydrogenase, but rather nitrogenase. Although beyond the scope of a review dedicated to hydrogenases, nitrogenase-mediated H2 production deserves mention on account of its global dimension. About 30–50 % of the total reducing power consumed by nitrogenase is side-tracked into the formation of H2 (Schubert and Evans 1976; Brewin 1984; Evans et al. 1987).

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H2-Metabolizing Prokaryotes

This phenomenon, a paradox considering the efficiency of other enzyme systems, raises the question of a biological role for this ‘‘side effect.’’ However, this may be, N2-fixing microbes often possess uptake hydrogenases which permit them to recover at least a part of the energy flowing into nitrogenase-mediated H2 production. Surprisingly, however, this is by no means a universal feature of diazotrophs. In many soil habitats, both H2-oxidizing and H2-nonoxidizing strains of the same diazotrophic species can be isolated. Therefore, significant quantities of H2 must escape into the environment (reviewed by Evans et al. 1987). This could be a major source of nutrient for H2-utilizing microbes such as the aerobic H2-oxidizing bacteria (La Favre and Focht 1983; Schuler and Conrad 1991b; Maimaiti et al. 2007). The capacity to reduce atmospheric dinitrogen is found both in archaea and bacteria, in aerobic as well as in anaerobic organisms (see > Chap. 11, ‘‘Dinitrogen-Fixing Prokaryotes’’ this volume for a list). Among the best-studied diazotrophs are the rhizobia including Bradyrhizobium japonicum, Sinorhizobium meliloti, and Rhizobium leguminosarum (reviewed in Hennecke 1990; Agron et al. 1994; Stacey et al. 1995; van Rhijn and Vanderleyden 1995; Ovtsyna et al. 2000; and Spaink 2000); strains of Azotobacter (reviewed by Peters et al. 1995); and various cyanobacteria (Fay 1992; Haselkorn and Buikema 1992). Many oxygenic and anoxygenic phototrophs fix dinitrogen and produce H2 concomitantly. Gest and Kamen discovered that cultures of R. rubrum grown in the light under nitrogenase-inducing conditions evolved significant amounts of H2 (Gest and Kamen 1949a, b). This process was referred to as ‘‘photoproduction’’ of H2. Subsequently, H2 photoproduction was observed in other phototrophic bacteria including R. capsulatus, R. gelatinosus, and R. palustris. Later investigations revealed that nitrogenase-catalyzed reduction of protons is the basis of H2 photoproduction (Bulen et al. 1965a, b). Recently, a different kind of photoproduction has been postulated. A hypothesis has been put forward that hydrogenases may act as redox buffers for the photosynthetic apparatus during transition from darkness to light (Appel and Schulz 1998; Appel et al. 2000). During such a transition, electrons from the photosynthetic apparatus could be channeled to H2 via a hydrogenase. Indeed, in Synechocystis, a darkness-to-light transition triggers a transient production of H2 (Abdel-Basset and Bader 1998). This hypothesis awaits confirmation by additional experimental studies.

Production of H2 as a Byproduct of Phosphite Oxidation Recently, a novel pathway for phosphite oxidation was discovered in E. coli (Yang and Metcalf 2004). The well-known periplasmic enzyme alkaline phosphatase (BAP) turned out to be the sole enzyme of this pathway, which catalyzes the following reaction: H3 PO3 þ H2 O ! H3 PO4 þ H2

ð4:6Þ

Thus, BAP is an H2-evolving hydrogenase. To date, BAP is the only microbial phosphatase show to have this activity.

H2-Consuming Processes Aerobic H2 Oxidation The aerobic hydrogen-oxidizing (knallgas) bacteria attracted the attention of microbiologists early on (Kaserer 1906; Niklewski 1910). These organisms utilize H2 as a source of energy via the oxyhydrogen reaction: 2H2 þ O2 ! 2H2 O

ð4:7Þ

The first part of the above net reaction, a heterolytic cleavage of H2, is catalyzed by various types of [NiFe] hydrogenases. It turned out that the first knallgas bacteria to be isolated and systematically studied were facultative H2 chemolithoautotrophs, which in the presence of sugars or organic acids grew organoheterotrophically. Furthermore, many of these organisms utilized H2 and organic substances mixotrophically (reviewed by Aragno and Schlegel 1992). The first obligate H2 chemolithotrophs were discovered much later. The aerobic thermophile Hydrogenobacter thermophilus is an obligate chemolithoautotroph (Kawasumi et al. 1984). The bacterium can, however, use H2 or elemental sulfur alternatively as electron donors. The marine bacterium Hydrogenovibrio marinus is an obligate H2 oxidizer (Nishihara et al. 1991, 1997). Mixotrophic H2 utilization probably plays an important role in pathogens of the human gastrointestinal tract. In the stomach pathogen Helicobacter pylori, for instance, hydrogenase significantly effects the efficiency of colonization of the gastric mucosa in the mouse experimental system (Olson and Maier 2002). Ralstonia eutropha (formerly Alcaligenes eutrophus) is one of the classical knallgas bacteria (Wilde 1962) and is now one of the best-studied H2 oxidizers. Ralstonia eutropha thrives on mixtures of H2 and CO2 but can, alternatively, utilize a broad spectrum of organic compounds. Moreover, it can also utilize H2 and organic substrates simultaneously. The bacterium contains two energy-generating [NiFe] hydrogenases: a membrane-bound type and a heterohexameric cytoplasmically localized species (> Fig. 4.2; Schneider and Schlegel 1976; Schink and Schlegel 1979). The membrane-bound enzyme is anchored to the periplasmic face of the cytoplasmic membrane and feeds electrons into a respiratory chain via a b-type cytochrome (> Fig. 4.2). This type of hydrogenase is very widespread and is the basis of H2 oxidation in most of the aerobic H2 oxidizers examined so far. The soluble, hexameric hydrogenase of R. eutropha was the first of its kind to be characterized genetically (Tran-Betcke et al. 1990). It consists of a hydrogenase moiety complexed with an NADH oxidoreductase (diaphorase) module (> Fig. 4.2). The enzyme couples the oxidation of H2 to the reduction of NAD+. A similar cytoplasmic [NiFe] hydrogenase is found in the Grampositive, facultative H2-oxidizer Rhodococcus opacus (formerly Nocardia opaca; Schneider et al. 1984a; Schneider et al. 1984b; Grzeszik et al. 1997b).

H2-Metabolizing Prokaryotes

Some diazotrophs are facultative chemolithoautotrophs. This dual strategy makes sense for inhabitants of the rhizosphere, since they often form hydrogenase to recycle H2 produced during N2 fixation and hence have the enzymatic tools to exploit H2 produced externally by other diazotrophs. The endosymbiotic N2-fixer Bradyrhizobium japonicum is an example for this group (Hanus et al. 1979). Outside the root nodule, B. japonicum is able to grow on H2 and CO2 as sole sources of energy and carbon. Azotobacter vinelandii is a free-living, strictly aerobic diazotroph. It is not an autotroph but thrives mixotrophically on H2 in the presence of organic substrates (Wong and Maier 1985). In the aerobic carboxydotrophs, CO can serve as the sole source of energy and carbon (reviewed in Meyer 1989). Many of these bacteria also thrive on H2 and CO2, and mixotrophic growth on H2 or CO in the presence of organic acids has also been reported (Kiessling and Meyer 1982). In Oligotropha carboxidovorans, oxidation of H2 is catalyzed by a membranebound hydrogenase (Santiago and Meyer 1997). Some phototrophs are facultative H2 chemolithotrophs (Bogorov 1974; Madigan and Gest 1979; Siefert and Pfennig 1979; Ka¨mpf and Pfennig 1980, 1986). This has been shown for Rhodobacter capsulatus, Rhodobacter sulfidophilus, and Rhodopseudomonas acidophila, which grow in the dark on H2 as sole source of energy and reducing power.

Methanogenesis Methanogenesis is one of the major H2-consuming processes in the biosphere. The global, annual rate of natural CH4 production has been estimated to be 190  70 Tg (Lelieveld et al. 1998). Hydrogenotrophic methanogens are true lithotrophs that convert H2 and CO2 to CH4 according to the following net reaction: 4H2 þ CO2 ! CH4 þ 2H2 O

ð4:8Þ

Other methanogens can utilize partially reduced forms of carbon such as methanol: CH3 OH þ H2 ! CH4 þ H2 O

ð4:9Þ

Whatever carbon compound is used as substrate, H2 is the source of reductant in most methanogens, and hence, hydrogenases participate at different stages of methanogenesis. The enzymatic pathway varies depending on the carbon substrate and organism. In general, there are two main types of methanogens: One group contains cytochromes and the other lacks them. Cytochromes are only found in members of the Methanosarcinales (reviewed in Thauer 1998; Keltjens and Vogels 1993; Ferry and Lessner 2008; Thauer et al. 2010). The production of CH4 from CO2 proceeds via an initial reaction in which N-formylmethanofuran is formed. In Methanosarcina barkeri, this reaction is dependent on reducing equivalents generated by a multisubunit hydrogenase (Ech) (Meuer et al. 2002; > Fig. 4.3). Similar membrane-bound complexes are thought to fulfill the same function in species of Methanothermobacter (Tersteegen and Hedderich 1999). Following the formation of

4

N-formylmethanofuran, the C1 unit is transferred to another carrier, tetrahydromethanopterin (H4MPT), and reduced in a stepwise fashion: N5, N10-methenyl-H4MPT is first converted to N5, N10-methylene-H4MPT. Subsequently, N5, N10-methyleneH4MPT is reduced to N5-methyl-H4MPT. A cytoplasmic [NiFe] hydrogenase, the heterotrimeric F420-reactive hydrogenase present in two isoenzymes, contributes reducing equivalents in the form of reduced cofactor F420 which drive these reactions (> Fig. 4.3). When deprived of nickel, M. thermoautotrophicus and some other methanogens produce high levels of an [Fe] hydrogenase, the so-called H2-forming H4MPT dehydrogenase (Hmd). Hmd is a cytoplasmic homodimer (> Fig. 4.3). This enzyme apparently substitutes for the nickel-containing F420reactive hydrogenase, catalyzing the reduction of coenzyme F420. In the final stage of methanogenesis, CH3-S-CoM and HS-CoB are oxidized to the heterodisulfide CoM-S-S-CoB liberating CH4 (> Fig. 4.3). The enzyme heterodisulfide reductase then regenerates the reduced forms of coenzyme M and coenzyme B via the reduction of CoM-S-S-CoB using reducing equivalents generated by hydrogenase. The heterodisulfide reductase differs from species to species. In strains of the Methanosarcinales, the enzyme is membrane bound. It receives its reducing power from a hydrogenase attached to the outer surface of the cytoplasmic membrane via the electron carrier methanophenazine (> Fig. 4.3). The reduction of CoM-S-S-CoB results in the formation of scalar protons and, hence, a gradient across the membrane which couples H2 oxidation to phosphorylation of ADP. In other species (e.g., strains of Methanothermobacter, Methanococcus and Methanopyrus), the heterodisulfide reductase and an F420-nonreducing hydrogenase form a soluble complex located in the cytoplasm (> Fig. 4.3; Setzke et al. 1994). Methanosarcina barkeri, M. mazei, and other methylotrophic methanogens can grow on substances such as methanol, methylamines, and acetate in addition to H2 and CO2 (Keltjens and Vogels 1993; Deppenmeier et al. 1999; Deppenmeier 2002). Regardless of what substrate is utilized, CH4 production leads to the formation of the heterodisulfide CoM-S-S-CoB. The pools of coenzymes HS-CoM and HS-CoB are replenished by the action of the heterodisulfide reductase. If adequate amounts of H2 are available, the reducing power for this reaction is supplied by membrane-bound methanophenazine-reducing hydrogenases (> Fig. 4.3). In the absence of H2, reducing power comes from a membrane-bound F420H2-dehydrogenase, which oxidizes coenzyme F420 in an energy-conserving manner. Both hydrogenase and F420H2-dehydrogenase are coupled to the heterodisulfide reductase via the redox intermediate methanophenazine (Deppenmeier 1995). The Ech hydrogenase of Methanosarcina barkeri is part of a multisubunit membrane-bound complex similar to hydrogenase-3 of E. coli (Ku¨nkel et al. 1998). Ech hydrogenase has multiple metabolic roles depending on the growth conditions (Meuer et al. 2002). During growth on H2 and CO2, Ech oxidizes H2, transferring the electrons to ferredoxin. Reduced ferredoxin provides the reducing power for the first step of methanogenesis, the reduction of CO2 to formylmethanofuran. In acetoclastic methanogenesis, Ech couples the oxidation of reduced

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H2-Metabolizing Prokaryotes

Ech hydrogenase (M. barkeri )

?

ΔµNa+ /ΔµH+

EchA Fe-S EchC EchB EchD

Fdox

[NiFe]

CHO-MFR

EchE

2H

+

5-CHO-H4MPT

[Fe] [Fe] [Fe] hydrogenase Hmd Hmd CH-H4MPT

[NiFe]

H2

VhoA

H+

F420

Fe-S b1 VhoG VhoC

CH2-H4MPT F420H2

2H+

FrhA Fe-S Fe-S FAD FrhG FrhB

b2 HdrE Fe-S HdrD

HS-CoB

HS-CoM

In

H2

Fdred Fdox

CH3-S-CoM HS-CoB

CH4

2H+

Heterodisulfide reductaseassociated hydrogenase (M. marburgensis)

CH3-H4MPT HS-CoM

CoM-S-S-CoB +2H+

Out

[NiFe]

F420

2H+

Heterodisulfide reductase

F420-reducing hydrogenase

F420H2

MPH2

2H+

MFR

H2

Methanophenazinereducing hydrogenase (M. mazei) H2

CO2

Fdred

Fe-S EchF

MP

140

CoM-S-S-CoB +2H+

Fe-S [NiFe] HdrC Fe-S MvhA Zn FAD Fe-S HdrA Fe-S HdrB MvhG

2H2 4H+

Fe-S MvhD

. Fig. 4.3 Archaeal hydrogenases: subunit composition, catalytic activity, and cellular location. MP, methanophenazine; MFR, methanofuran; CHOMFR, formylmethanofuran; HS-CoB, coenzyme B; HS-CoM, coenzyme M; Fdred/Fdox, reduced/oxidized ferredoxin; b1 and b2 cytochromes, b1 and b2, respectively. See text for additional details

ferredoxin (arising from the oxidation of the carbonyl group of acetate) to the production of H2. Finally, during growth on methanol, H2, and CO2, Ech seems to have a biosynthetic role (Deppenmeier 2002; Hedderich and Forzi 2005). Methanosaeta thermophila is an interesting exception to the above paradigms. This obligate aceticlastic methanogen lacks membrane-bound hydrogenases including Ech-type complexes (Smith and Ingram-Smith 2007; Welte and Deppenmeier 2011). The energy-conserving mechanisms of this organism must differ fundamentally from other Methanosarcinales.

The reducing power for this reaction is ultimately derived from H2 by the action of hydrogenases (Drake 1982; Pezacka and Wood 1984). Acetobacterium woodii contains a soluble hydrogenase of the [FeFe] type (Ragsdale and Ljungdahl 1984). Sporomusa sphaeroides oxidizes H2 with the help of a heterodimeric, membrane-bound [NiFe] hydrogenase (Dobrindt and Blaut 1996).

Sulfate and Sulfur Reduction

When organisms such as Acetobacterium woodii, Moorella thermoacetica, Moorella thermoautotrophica, and Clostridium aceticum grow on CO2 and H2, CO2 is converted to acetate via the Ljungdahl-Wood pathway (Diekert and Wohlfarth 1994). This reaction proceeds via the following stoichiometry:

The different strains of sulfate-reducing bacteria and archaea use a spectrum of electron donors including alcohols, H2, and organic acids such as acetate, lactate, malate, and pyruvate (reviewed by > Chap. 9, ‘‘Dissimilatory Sulfate- and SulfurReducing Prokaryotes’’ this volume). The electron acceptor, sulfate, sulfite, thiosulfate, or elemental sulfur, is reduced to sulfide via the following net reactions: ð4:11Þ 4H2 þ H2 SO4 ! H2 S þ 4H2 O

2CO2 þ 4H2 ! CH3 COOH þ 2H2 O

3H2 þ H2 SO3 ! H2 S þ 3H2 O

Acetogenesis

ð4:10Þ

ð4:12Þ

H2-Metabolizing Prokaryotes

4H2 þ H2 S2 O3 ! 2H2 S þ 3H2 O

ð4:13Þ

H2 þ S ! H2 S

ð4:14Þ

Thus, the oxidation of an organic compound or H2 is coupled to the reduction of a sulfur compound. This process entails the generation of a proton gradient across the cytoplasmic membrane. Energy is conserved via a chemiosmotic mechanism, for which reason the process is loosely called ‘‘anaerobic respiration’’ (Brandis and Thauer 1981). Many sulfate reducers thrive on H2 as the sole source of energy and reductant. This group includes strains of Desulfovibrio, Desulfomicrobium, Desulfobacter, Desulfobacterium, Desulfonema, Desulfobulbus, Desulfosarcina, Thermodesulfovibrio, and Thermodesulfobacterium (see > Chap. 9, ‘‘Dissimilatory Sulfate- and SulfurReducing Prokaryotes’’ this volume). Several of these strains can grow lithoautotrophically on H2 and CO2 (Jansen et al. 1984; Moller et al. 1987; Schauder et al. 1989). Autotrophic growth on H2 and CO2 has been documented for Desulfovibrio fructosovorans, Desulfomicrobium apsheronum (Rozanova et al. 1988), Desulfobacterium autotrophicum (Brysch et al. 1987), Desulfotomaculum geothermicum (Daumas et al. 1988), Desulfotomaculum kuznetsovii (Rozanova et al. 1988), Desulfobacter hydrogenophilus, Desulfomonas limicola, and Desulfosarcina variabilis (Widdel 1988). The genome sequence of a sulfate-reducing organism has been assembled from DNA samples collected in an aquifer at a depth of 2.8 km. Based on the genomic data, an autotrophic, H2-oxidizing, N2-fixing sulfate reducer named ‘‘Desulforudis audaxviator’’ was postulated (Chivian et al. 2008). Perhaps the most remarkable property of sulfate reducers is their mixotrophic metabolism, that is, their ability to utilize H2 and organic compounds simultaneously. In this case, energy is generated by two different modes operating at the same time: (1) electron-transport phosphorylation driven by the hydrogenase-dependent respiratory chain and (2) substrate-level phosphorylation coupled to the oxidation of an organic substrate. Hydrogen is one of the products of the latter process. If sufficient sulfate is present, net production of H2 seldom occurs. Hydrogen produced via fermentation is reoxidized via the hydrogenase-dependent respiratory chain (Tsuji and Yagi 1980). Under low-sulfate conditions, H2 is produced and released into the environment (Postgate 1952; Vosjan 1975; Hatchikian et al. 1976; Traore et al. 1981). Extensive biochemical and genetic studies on sulfatereducing bacteria have revealed the existence of multiple hydrogenases in one and the same organism (Voordouw 1992). As genome sequences become available, it is possible to predict the inventory of hydrogenase for an increasing number of organisms (Baltazar et al. 2011). Desulfovibrio vulgaris Hildenborough, for instance, carries genes for 7 different hydrogenases (Heidelberg et al. 2004). Of these, 4 are periplasmically oriented enzymes (3 [NiFe]-type and 1 [FeFe]-type). The other 3 are cytoplasmically oriented (2 [NiFe]-type and 1 [FeFe]-type). Syntrophobacter

4

fumaroxidans MPOB has genetic determinants for nine different hydrogenases (de Bok et al. 2002; Baltazar et al. 2011). The assignment of physiological functions to multiple hydrogenases is difficult. One approach to the problem is the analysis of mutants with defined genetic lesions. Desulfovibrio fructosovorans forms at least four different hydrogenases (Casalot et al. 2002a): Two of these, an [FeFe] and a [NiFe] enzyme, are periplasmic. The third hydrogenase is a heterotetrameric, cytoplasmic, NADP-dependent [FeFe] enzyme (Rousset et al. 1990; Malki et al. 1995, 1997). The former enzymes are probably part of an H2-dependent respiratory chain. The latter could couple H2 oxidation directly to the generation of reducing equivalents. Mutants defective for the [NiFe] hydrogenase or the cytoplasmic hydrogenase or both still grew well on H2 and sulfate. During growth on fructose, lactate, and pyruvate, the mutants behaved differently: The strain defective for the [NiFe] enzyme grew as well as the wild type. Growth of the strains lacking a functional cytoplasmic hydrogenase was significantly curtailed (Malki et al. 1997). Thus, the bioenergetic contributions of the three hydrogenases are not sharply defined and seem to allow for a certain degree of mutual compensation. However, the cytoplasmic [FeFe] hydrogenase is evidently more important during growth involving fermentative utilization of organic substrates. A triple mutant defective for all three of the hydrogenases described above grew at a rate similar to the wild type on H2 (Casalot et al. 2002b). This suggests that there is at least one additional hydrogenase in D. fructosovorans that has not been identified. The four periplasmically oriented hydrogenases of Desulfovibrio vulgaris Hildenborough comprise three soluble hydrogenases (encoded by the hyn1, hys, and hyd genes) and a membrane-attached enzyme (encoded by hyn genes). HysAB is a [NiFe(Se)] hydrogenase, HydAB is an [FeFe] hydrogenase, and the two Hyn enzymes contain [NiFe]. A Desulfovibrio vulgaris mutant defective for the cytoplasmic [FeFe] hydrogenase showed reduced growth both on H2 and on lactate (Pohorelic et al. 2002). Under the latter conditions, the mutant also liberated much more H2 than the wild type. Thus, the Hyd hydrogenase is involved in the utilization both of external H2 and of internally generated H2 arising from the fermentation of organic substrates. hyn1 and hys mutants were only slightly affected in growth on lactate or high levels of H2 (50 % v/v). However, the hyn1 mutant stopped growing at a lower cell density than the wild type, and survival in the stationary phase was curtailed (Goenka et al. 2005). At lower H2 levels (5 % v/v), growth of the hys mutant was significantly impaired. This suggests that the organism produces hydrogenases with different Km values as a way of adapting to fluctuating concentrations of environmental H2 (Caffrey et al. 2007). The respiration of sulfate with H2 as electron donor has been reported for other groups of bacteria aside from the classical sulfate reducers. One example of this is Allochromatium minutissimum (Nakamura 1939, 1941), which grows anaerobically in the dark on H2 and sulfate. The existence of multiple hydrogenases of different types in many sulfate reducers underscores their important metabolic role in these organisms. Nevertheless, genome projects have led to the surprising discovery that some sulfate reducers lack hydrogenases altogether.

141

142

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H2-Metabolizing Prokaryotes

The archaeon Caldivirga maquilingensis is an example. Evidently, sulfate respiration is not necessarily linked to H2 oxidation. In some anaerobic H2-oxidizers, sulfur takes the place of sulfate as electron acceptor in the oxidation of H2 (see Hedderich et al. 1999 for a list). These organisms are typically found in hot, marine solfataras. The marine thermophilic archaeon Thermoproteus neutrophilus is an obligate chemolithoautotroph, growing on H2 and elemental sulfur with CO2 as sole carbon source (Fischer et al. 1983). Its relatives, Thermoproteus tenax and Pyrobaculum islandicum, are H2-utilizing chemolithotrophs that can grow in a facultative chemoorganotrophic mode, respiring sulfur (Fischer et al. 1983). Pyrodictium occultum and Pyrodictium brockii are both hyperthermophilic archaea that obtain energy via H2/S chemolithotrophy (Fischer et al. 1983; Stetter and Gaag 1983). In the latter organism, H2 oxidation is catalyzed by a heterodimeric, membrane-bound [NiFe] hydrogenase (Pihl et al. 1989; Pihl and Maier 1991). The moderately thermophilic, strictly anaerobic bacterium Desulfurella multipotens is a facultative chemolithotroph growing on H2 and elemental sulfur (Miroshnichenko et al. 1994). It has now been recognized that sulfur-metabolizing Epsilonbacteria make up a significant part of the biodiversity in hydrothermal settings (Campbell et al. 2006). Organisms such as Nautilia lithotrophica and Caminibacter mediatlanticus oxidize H2 using sulfur compounds or nitrate as electron acceptors (Miroshnichenko et al. 2002; Voordeckers et al. 2005). Mixotrophic growth has been reported for another member of the genus Pyrodictium: the marine hyperthermophile Pyrodictium abyssi (Pley et al. 1991). Unlike the other two species, it is a heterotroph, growing by fermentation of organic compounds. The addition of elemental sulfur to heterotrophic cultures has little effect on growth. However, when H2 and S are provided in addition to organic substrates, growth is markedly stimulated and sulfur is reduced to H2S. Perhaps, the most impressive examples of the metabolic versatility of the sulfur-reducers are the archaeon Acidianus infernus and its relatives (Segerer et al. 1986). Under anaerobic conditions, these organisms grow by H2/S chemolithotrophy. They are, however, facultative aerobes: In the presence of O2, they oxidize sulfur to sulfuric acid. The exploitation of a variety of electron acceptors for the oxidation of H2 is not limited to archaea. The bacterium Sulfurospirillum deleyianum (Schumacher et al. 1992; Eisenmann et al. 1995) can grow on H2 or elemental sulfur as sole source of energy and reducing power. It utilizes a palette of electron acceptors including oxygen, nitrate, nitrite, sulfur compounds, and organic acids. The hyperthermophile Aquifex pyrophilus (Huber et al. 1992) grows on H2 in the presence of O2 just like a conventional knallgas bacterium. Alternatively, both S0 and S2O3 2 can serve as electron donors in place of H2. Under anoxic conditions, A. pyrophilus can grow on H2, S0, or S2O3 2 using nitrate as the terminal electron acceptor. Furthermore, in the late stage of exponential growth on H2 under oxic conditions, A. pyrophilus switches from O2 to S0 as the terminal electron acceptor and begins producing H2S.

The model organism for biochemical and molecular studies on anaerobic respiration is Wolinella succinogenes (Hedderich et al. 1999; Lancaster 2001). This anaerobic, rumen organism oxidizes H2 using sulfur in the form of polysulfide as the terminal electron acceptor (Jacobs and Wolin 1963; Kro¨ger and Innerhofer 1976; Bronder et al. 1982; Jankielewicz et al. 1995). The membrane-bound [NiFe] hydrogenase is coupled to the membrane-bound polysulfide reductase via the redox carrier menaquinone (Dross et al. 1992).

Fe(III) Reduction Many strains of Fe(III)-reducing bacteria are capable of using H2 as an electron donor. The coupling of H2 oxidation to the reduction of Fe(III) is an important energy-yielding process in subsurface microbial communities. Members of the family Geobacteraceae are also capable of reducing other metals such as Mn(IV) and U(VI). Both Geobacter hydrogenophilus and Geobacter sulfurreducens grow on H2 (Coates et al. 2001; Caccavo et al. 1994). In Geobacter sulfurreducens, H2 oxidation is catalyzed by a membrane-bound hydrogenase (Coppi et al. 2004). H2-based lithotrophic growth with Fe(III) as terminal electron acceptor has been described in both archaeal thermophiles (e.g., Pyrobaculum islandicum and Thermotoga maritima), suggesting that H2-oxidation with Fe(III) is one of the earliest forms of respiration (Vargas et al. 1998; Kashefi and Lovley 2000).

Dehalorespiration Certain bacteria exploit specialized respiratory chains, in which the oxidation of H2 or organic acids is coupled to the dehalogenation of haloaliphatic or haloaromatic compounds. This type of energy metabolism is known as dehalorespiration and has been found in both Gram-negative and Gram-positive bacteria. Hydrogenases and reductive dehalogenases are key components of these pathways. Among the representatives of this group are both strict H2 oxidizers, such as Dehalobacter restrictus (Holliger et al. 1998), and organisms, such as Dehalospirillum multivorans (Scholz-Muramatsu et al. 1995), which can utilize a variety of electron donors. Both of these strains contain membrane-bound hydrogenases. Dehalococcoides ethenogenes is another strict H2 oxidizer. The genome sequence of this organism reveals the genetic determinants for five different hydrogenases (Seshadri et al. 2005).

Anoxygenic Photosynthesis In the versatile metabolism of the anoxygenic phototrophic bacteria, H2 has different roles depending on the growth conditions (Vignais et al. 1985). Two of the metabolic functions of H2 have been discussed above: H2 is consumed as a source of energy and reductant during aerobic, chemolithoautotrophic growth and is produced as a fermentation product during anaerobic,

H2-Metabolizing Prokaryotes

heterotrophic growth. A third role of H2 is linked to anaerobic growth in the light: Many anoxygenic phototrophs can utilize H2 as an electron donor for photoautotrophic growth. This was first shown for the purple sulfur bacterium Allochromatium minutissimum (Roelofsen 1934; Gaffron 1935). Photosynthetic H2 oxidation is dependent on uptake hydrogenases (Gest 1951). Both purple nonsulfur and purple sulfur bacteria are capable of photosynthetic growth on H2 (see Drews and Imhoff 1991 for a list). Included in this group are Rhodospirillum rubrum (Ormerod and Gest 1962; Anderson and Fuller 1967), Rhodopseudomonas palustris (Qadri and Hoare 1968), Rubrivivax gelatinosus (Wertlieb and Vishniac 1967), Rhodobacter capsulatus (Klemme and Schlegel 1967), Allochromatium vinosum (Gitlitz and Krasna 1975), and Thiocapsa roseopersicina (Gogotov 1968). In some if not all of the above-named organisms, oxidation of H2 for photosynthetic growth is catalyzed by membrane-bound [NiFe] hydrogenases (Gitlitz and Krasna 1975; Adams and Hall 1977; Bagyinka et al. 1982; Kondratieva and Gogotov 1983; Kovacs et al. 1983; Gogotov 1984; Zorin et al. 1996b; > Fig. 4.2). The marine sulfur bacterium Thiocapsa roseopersicina BBS contains four [NiFe] hydrogenases (Colbeau et al. 1994; Ra´khely et al. 1998, 2004; Kova´cs et al. 2002, 2005c; Maro´ti et al. 2010). Two of these hydrogenases are membrane-bound enzymes; two others are located in the cytoplasmic. The four enzymes have different but overlapping profiles of H2-evolving and H2consuming activity indicating a certain degree of physiological specialization (Ra´khely et al. 2007). The heteropentameric Hox1 enzyme is responsible for H2 evolution during fermentative growth in the dark, thus maintaining the redox balance of the cell. However, Hox1 also contributes to H2 oxidation during phototrophic growth. The Hup hydrogenase oxidizes H2 in the dark under aerobic conditions and is probably the basis for the chemolithotrophic growth mode. Hup is also active during phototrophic, anaerobic growth. Under these conditions, its role may be to supply electrons for photosynthesis. Less clear is the role of Hox2. Hox2 has both H2-evolving and H2-oxidizing activity in vivo, but its contribution seems to be minor compared to the other hydrogenases (Maro´ti et al. 2010). Under N2-fixing conditions, the H2-oxidizing hydrogenases recycle H2 produced by nitrogenase. This is probably one of the functions of the Hyn hydrogenase (> Fig. 4.2).

Fumarate Respiration E. coli produces two membrane-bound [NiFe] uptake hydrogenases under anaerobic conditions (overview in Forzi and Sawers 2007). Studies using defined mutants clarify the physiological specialization of these isoenzymes. Hydrogenase-2 (Hyd-2) supports growth on H2 and fumarate as sole sources of energy and carbon, respectively. Hyd-2 deficient mutants do not grow under these conditions, whereas the growth of hydrogenase-1 (Hyd-1) knockout mutants is not impaired (Dubini et al. 2002). Hyd-2 is a respiratory enzyme which couples the oxidation of external H2 to the reduction of

4

fumarate inside the cell, thereby contributing to the generation of proton motive force. The synthesis of Hyd-1 is stimulated by carbon limitation, suggesting a role for this enzyme in stationary phase survival (Pinske et al. 2012).

Ancillary Processes This category comprises H2-consuming metabolic activities that are allied to and dependent upon H2-evolving processes in one and the same cell. Here, we are dealing with hydrogenases whose primary purpose is to consume internally produced H2. Hydrogenase-free strains of the organisms in question are readily isolated from natural habitats and thrive despite the lack of H2-activating enzymes. The foremost activity of this kind is the so-called hydrogen recycling observed in many diazotrophs. This is the role of the heterodimeric, membrane-bound [NiFe] hydrogenases in the endosymbiotic N2-fixers such as Sinorhizobium leguminosarum (Brewin 1984). A similar function can be ascribed to free-living N2-fixers such as Azotobacter chroococcum (Ford et al. 1990), Bradyrhizobium japonicum (Harker et al. 1984; Sayavedra-Soto et al. 1988), Rhodobacter capsulatus (Willison et al. 1983; Leclerc et al. 1988), and Rhodocyclus gelatinosus (Uffen et al. 1990). In the latter two cases, however, hydrogenases can also catalyze the oxidation of externally available H2 and hence support facultative chemolithotrophic growth. Another example which falls into this category is the hydrogenase-mediated consumption of H2 which is evolved as a terminal product of fermentation. Desulfovibrio vulgaris and other sulfate reducers evolve H2 both during growth on SO4 2 and during fermentation. A portion of this H2 is reoxidized by hydrogenase in an energy-conserving mechanism called ‘‘hydrogen cycling’’ (Odom and Peck 1981; Lupton et al. 1984). The methanogen M. barkeri deserves mention in this context. During growth on acetate, H2 is produced by the Ech hydrogenase (Meuer et al. 2002). The internally produced H2 is consumed by the methanophenazine-reducing hydrogenases (> Fig. 4.3).

Classification of Hydrogenases The first hydrogenases to be isolated and characterized biochemically were the iron-containing enzymes of sulfur-reducing bacteria. Later on, it was discovered that some hydrogenases contain nickel in addition to iron (Friedrich et al. 1981b; Graf and Thauer 1981). The preponderance of hydrogenases which have been characterized to date is of this type. Some of these enzymes contain selenium in the form of the unusual amino acid selenocysteine (Rieder et al. 1984; He et al. 1989). Finally, [Fe] hydrogenases have been found in certain methanogens (Thauer et al. 1996). These enzymes, called ‘‘H2-evolving N5, N10-methylenetetrahydromethanopterin dehydrogenases,’’ or ‘‘Hmd hydrogenases,’’ are neither mechanistically nor structurally related to other hydrogenases. Originally classified as ‘‘metal-free hydrogenases,’’ the Hmd’s were subsequently

143

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H2-Metabolizing Prokaryotes

shown to contain iron. Unlike the classical iron hydrogenases, however, the Hmd’s harbor a cofactor containing a single iron atom, but no iron-sulfur centers (Lyon et al. 2004a, b). Early classification schemes, which had mainly biochemical data to go on, grouped hydrogenases on the basis of metal content or redox cofactors (Fauque et al. 1988; Przybyla et al. 1992). A rapidly growing base of nucleotide sequence data prompted Voordouw to attempt a classification on the basis of comparisons of deduced amino acid sequences (Voordouw 1992). In a similar study, Wu and Mandrand (1993) went a step further. These authors generated multiple alignments for full-length amino acid sequences and performed cluster analysis on the pairwise alignment scores. On the basis of the resulting dendrograms, 30 hydrogenases were grouped in six classes. Recently, Vignais and coworkers have refined and extended this classification system, including a greatly expanded database (Vignais et al. 2001; Vignais and Billoud 2007). They carried out thorough cluster analyses using both complete amino acid sequences and segments corresponding to functional domains. The results of these important studies support the notion that hydrogenases belong to three phylogenetically distinct groups: [FeFe] hydrogenases, [NiFe] hydrogenases (including [NiFe(Se)] hydrogenases), and [Fe] hydrogenases. Thus, the original, pragmatic classification now has a foundation in molecular phylogenetics.

[NiFe] Hydrogenases The revised system of Vignais and coworkers subdivides the [NiFe] hydrogenases into four groups (Vignais et al. 2001; Vignais and Billoud 2007): Group 1. Energy-Transducing Hydrogenases

These are enzymes which couple the oxidation of H2 to electrontransport phosphorylation. This group includes both membrane-bound hydrogenases attached to the periplasmic side of the cytoplasmic membrane and nonmembrane-bound, periplasmic hydrogenases. The group breaks down into two subclusters. One contains the membrane-bound hydrogenases of the proteobacterial type and the other, archaeal membrane-bound hydrogenases. Group 2. Cytoplasmic Uptake Hydrogenases

Group 2a contains cyanobacterial soluble uptake hydrogenases. Group 2b contains sensory hydrogenases. These soluble proteins are components of signal-transmitting circuits governing the expression of hydrogenase and accessory genes. Specialized hydrogenases of this type have to date been studied in detail in Bradyrhizobium japonicum and strains of Ralstonia and Rhodobacter. Group 3. Multimeric Cytoplasmic Hydrogenases

These are enzymes of complex subunit composition, which interact with soluble cofactors. The group breaks down into four subgroups consisting of F420-reducing

hydrogenases of methanogenic archaea (group 3a), NADPreducing hydrogenases of bacteria and archaea (group 3b), methyl viologen-reducing hydrogenases of archaea (group 3c), and the so-called bidirectional hydrogenases of R. eutropha, R. opacus, and cyanobacteria (group 3d). Group 4. Multisubunit, Membrane-Bound Energy-Converting Hydrogenases

The best-studied representative is E. coli hydrogenase-3, an H2-evolving enzyme, which is part of the multisubunit formate-hydrogenlyase complex. Similar enzymes have been identified in various bacteria and archaea including R. rubrum, C. hydrogenoformans, M. barkeri, M. thermoautotrophicus, and P. furiosus. Group 5. High-Affinity Hydrogenases

A relatively recent extension of the classification scheme is a fifth group comprising putative high-affinity hydrogenases (Constant et al. 2011). This group includes hydrogenases identified in Streptomyces avermitilis and many other Actinobacteria but also in a wide variety of taxa belonging to Proteobacteria, Chloroflexi, and Acidobacteria.

[FeFe] Hydrogenases The [FeFe] hydrogenases are quite heterogeneous in quaternary structure and domain organization (Vignais et al. 2001). Monomeric, dimeric, trimeric, and tetrameric enzymes have been described. Moreover, the basic hydrogenase domain is coupled to various other functional modules in one and the same subunit. Even analyses confined to partial sequences corresponding to the conserved hydrogenase catalytic site (H-cluster) fail to show well-separated subgroups (Vignais et al. 2001). Thus, a subdivision of the [FeFe] hydrogenases is not feasible at present.

[Fe] Hydrogenases A third class of hydrogen-activating enzymes consists of the Hmd’s. Originally discovered in Methanothermobacter marburgensis, Hmd enzymes have been found in other methanogenic archaea including Methanococcus voltae and Methanopyrus kandleri (Thauer et al. 1996, 2010). The extensive sequence identity in this class of enzymes shows that they are highly conserved.

Biochemistry of the Hydrogen-Converting Enzymes The old observation that knallgas bacteria require nickel for growth on H2, O2, and CO2 (Bartha and Ordal 1965; Tabillion et al. 1980) took on new significance when it was noticed that nickel is essential for the biosynthesis of active

H2-Metabolizing Prokaryotes

CN

CO

3Cys[4Fe-4S]

Cys

Cys

[Fe] hydrogenase

[FeFe] hydrogenase

[NiFe] hydrogenase

Cys

Bridgehead atom Fe

OH−

Fe2 Cys

CN

Fe1

CN

Cys CO

CO

Fe

Cys CO

GMP Pyridol

H2O

Ni

4

CO

CN

H 2O

CO

. Fig. 4.4 The catalytic sites of the three classes of hydrogenases as deduced from X-ray crystallography. Metal atoms and coordinating water/ hydroxide molecules are shown as spheres. Nonmetal ligands are represented as sticks (yellow: sulfur; gray: carbon; red: oxygen; blue: nitrogen). The [NiFe] site of the periplasmic hydrogenase of Desulfovibrio gigas is depicted (Volbeda et al. 1996; PDB entry: 2FRV). The Hcluster of the [FeFe] hydrogenase of Clostridium pasteurianum is shown (Pandey et al. 2008; PDB entry: 3C8Y); the central atom of the bridgehead is drawn as a nitrogen following Silakov et al. (2009), and the mononuclear iron site of the [Fe] hydrogenase of Methanocaldococcus jannaschii is displayed (Hiromoto et al. 2009a; PDB entry: 3F47).

hydrogenase in Ralstonia eutropha (Friedrich et al. 1981b) and it was discovered at the same time that hydrogenase purified from Methanothermobacter marburgensis contains this transition metal in stoichiometric amounts (Graf and Thauer 1981). Since then, extensive genetic, biochemical, and spectroscopic analyses have shed light on the molecular structure and catalytic mechanism of the three well-known hydrogenase classes: [NiFe], [FeFe], and [Fe] hydrogenases. These phylogenetically unrelated groups of hydrogenases share a complex active site that consists of at least one iron coordinated by varying numbers of cysteine residues and biologically unique carbon monoxide (CO) and cyanide (CN ) ligands (> Fig. 4.4). This section of the survey focuses on the structure and function of the catalytic sites and their interaction with various prosthetic groups either embedded in the same polypeptide or located on additional subunits. The scarcely understood H2 production activity of nitrogenase (reviewed by Seefeldt et al. 2012) and of the nickel-containing CO dehydrogenase II from Carboxydothermus hydrogenoformans (Svetlitchnyi et al. 2001) is not addressed in this article. Likewise, the recently discovered H2 oxidation activity of two energyconverting selenium- and molybdenum-dependent formate dehydrogenases from Escherichia coli (Soboh et al. 2011) is not discussed.

[NiFe] Hydrogenases The Basic Module The basic module of catalytically active [NiFe] hydrogenase consists of two heterologous subunits of approximately 40–70 and 16–30 kDa that share sequence similarity with domains of the NuoD and NuoB subunits of complex I in mitochondria (NADH-ubiquinone oxidoreductase; reviewed by FontecillaCamps et al. 2007; Vignais and Billoud 2007). Crystal structures

of several [NiFe] hydrogenases (Volbeda et al. 1995; Higuchi et al. 1997; Montet et al. 1997; Matias et al. 2001) and of two [NiFe(Se)] hydrogenases (Garcin et al. 1999; Marques et al. 2010) from anaerobic sulfate-reducing bacteria have been solved. Recently, this was also accomplished for two O2-tolerant [NiFe] hydrogenases isolated from knallgas bacteria (Fritsch et al. 2011b; Shomura et al. 2011) and for hydrogenase-1 of E. coli (Volbeda et al. 2012). In the following, we describe the structure of the prototypic [NiFe] hydrogenase of Desulfovibrio gigas in detail and subsequently compare it to other representatives of this class of enzymes. The periplasmic [NiFe] hydrogenase of D. gigas contains three Fe-S clusters in the small subunit, whereas the [NiFe]-containing active site is located in the large subunit (Hatchikian et al. 1978; Cammack et al. 1982; Huynh et al. 1987), coordinated by an N-terminal and a C-terminal pair of cysteines (> Fig. 4.5). Two of the thiolate groups (provided by Cys68 and Cys533) form a bridge between the two metals (Volbeda et al. 1995). In the case of [NiFe(Se)] hydrogenases, the nickel-ligating Cys530 is replaced by a selenocysteine residue (Garcin et al. 1999; Marques et al. 2010). Electron density maps of the oxidized, catalytically inactive [NiFe] hydrogenase from sulfate-reducing bacteria identified hydroxide and peroxide species—the latter is still a matter of debate—as bridging ligands between the Ni and the Fe atom. The bridging ligands disappear upon reduction, a process which correlates with the onset of the catalytic cycle (reviewed by Ogata et al. 2009). Three further characteristic electron densities were observed at the Fe atom. Fourier transform infrared (FTIR) spectroscopy, first applied to the [NiFe] hydrogenase of Allochromatium vinosum (formerly Chromatium vinosum), uncovered three distinct infrared bands in the high stretching frequency region (Bagley et al. 1994, 1995). These bands could later be assigned to three diatomic nonprotein ligands, one CO and two CN molecules, bound to the Fe at the heterodinuclear site

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H2-Metabolizing Prokaryotes

Cys530L

Fe Ni Cys68L

Cys533L

Cys65L

10.7 Å Cys17S Cys20S

[4Fe-4S] proximal [3Fe-4S] medial 9.5 Å Cys148S

Cys228S

[4Fe-4S] distal

Cys112S Cys246S Cys249S

Cys188S

8.6 Å Cys219S Cys213S His185S

. Fig. 4.5 The three-dimensional structure of the D. gigas [NiFe] hydrogenase (PDB entry: 2FRV). The plane at the interface between the two subunits is indicated by a thick, dashed line. The large subunit (to the left and above the gray bar) harbors the active site. The small subunit coordinates three iron-sulfur clusters (red: iron; yellow: sulfur). Cysteine residues, which are involved in the coordination of the active-site metals, are labeled. Xenon atoms (blue spheres) were used to probe for a gas tunnel in the similar protein from Desulfovibrio fructosovorans (Montet et al. 1997). The blue grid indicates a putative gas tunnel that connects the catalytic site with the external medium. The blow-up shows the three diatomic ligands and the bridging ligand (X). The latter varies depending on the redox state of the [NiFe] site (Adapted from Frey et al. (2000) and Frey et al. (2001))

(Happe et al. 1997; Pierik et al. 1999). FTIR studies on the D. gigas [NiFe] hydrogenase confirmed the assignment of one CO and two CN ligands (> Fig. 4.5). Modeling predicts that the two CN ligands accept hydrogen bonds from the amino acids Arg463 and Ser486, whereas the CO ligand is surrounded by hydrophobic residues (Volbeda et al. 1995, 1996). However, the exact orientation of the three ligands remains to be elucidated. The standard type of [NiFe] hydrogenase forms a globular heterodimer with a radius of approximately 60 A˚. The two subunits share a large planar surface (> Fig. 4.5) stabilized via ionic

e-

. Fig. 4.6 Postulated electron transfer pathway in the [NiFe] hydrogenase of D. gigas (PDB entry: 2FRV). The [NiFe] active site is shown along with the Fe-S centers and ligating amino acid side chains (yellow: sulfur; gray: carbon; red: oxygen; blue: nitrogen; brown: iron). Key distances between metal atoms are labeled and shown as dashed lines. Residues are labeled according to their location in the large (L) and small (S) hydrogenase subunits

interactions (Szilagyi et al. 2002). The catalytic site and the proximal [4Fe-4S] cluster are buried deep within the protein (Volbeda et al. 1995). Spectroscopic data and the spatial arrangement of the proximal [4Fe-4S], the medial [3Fe-4S], and the distal [4Fe-4S] cluster in the small subunit, separated by intervals of 8.6–10.7 A˚, suggest that the Fe-S clusters function as a ‘‘conductive wire.’’ The proximal [4Fe-4S] cluster can exchange electrons directly with the catalytic site (> Fig. 4.6). The N-terminal part of the electron-transferring small subunit contains a highly conserved flavodoxin-like domain providing the binding site for the proximal [4Fe-4S] cluster. This is obviously an essential feature of all [NiFe] hydrogenases. The role of the medial [3Fe-4S] center in electron transfer is still a matter of debate, since its redox potential seems to be too high relative to that of the H2 reactive site (Albracht 2001; Fontecilla-Camps et al. 2001). The distal [4Fe-4S] cluster of the D. gigas hydrogenase is coordinated by three cysteines and one histidine residue. This solvent-exposed histidine appears to be indispensible for the electronic exchange between the hydrogenase and its corresponding redox partner, for example, a multiheme cytochrome c3 (Dementin et al. 2006). In the periplasmic [NiFe(Se)] hydrogenases, the set of amino acids responsible for coordinating the medial Fe-S center contains a cysteine at the position of the conserved proline residue.

H2-Metabolizing Prokaryotes

Indeed, a medial [4Fe-4S] cluster was found in the crystal structure of the Desulfomicrobium baculatum and Desulfovibrio vulgaris Hildenborough hydrogenases (Garcin et al. 1999; Marques et al. 2010). A similar Fe-S cluster composition has been reported for the [NiFe(Se)] hydrogenase of Methanococcus voltae, and conversion of the medial [4Fe-4] center to a [3Fe-4S] form by site-directed mutagenesis yielded a protein having near wild-type activity with benzyl viologen but drastically decreased activity with the physiological acceptor coenzyme F420 (Bingemann and Klein 2000). The conversion of the native [3Fe-4S] to a [4Fe-4S] cluster in the [NiFe] hydrogenase of D. fructosovorans had little effect on the enzymatic activity irrespective of whether redox dyes or cytochrome c3 were used as electron acceptors. Interestingly, the mutation changed the ratio of H2 uptake to H2 production activity significantly and conferred increased oxygen sensitivity to the mutant protein (Rousset et al. 1998b). This observation supports the notion that in some organisms, the [3Fe-4S] cluster protects the enzyme from inactivation by oxygen (Albracht 1994). Based on amino acid sequence analyses, a medial [4Fe-4S] center was postulated for the H2-sensing [NiFe] hydrogenases (Kleihues et al. 2000). A few [NiFe] hydrogenases, including the NAD-reducing multimeric hydrogenases and the E. coli hydrogenase-3, contain a minimal version of a functional hydrogenase module consisting of a [NiFe] active site in the large subunit and a single proximal [4Fe-4S] cluster in the small subunit. It was demonstrated with R. eutropha and E. coli mutants that this hydrogenase module represents the minimal functional unit required for catalytic activity (Massanz et al. 1998; Pinske and Sawers 2011).

The [NiFe] Active Site Comparison of a large number of deduced amino acid sequences clearly shows that [NiFe] hydrogenases share common motifs. The [NiFe] cofactor-harboring subunit contains a set of at least five related signatures in the N-terminal and C-terminal regions that are located close to the active site (Voordouw et al. 1989; Wu and Mandrand 1993). A systematic site-directed mutagenesis study conducted with the multimeric NAD-reducing hydrogenase of R. eutropha shed light on the possible role of these conserved residues in the catalytic cycle (Massanz and Friedrich 1999; Burgdorf et al. 2002). The pattern, inferred from multiple sequence alignments of two specific motifs, the so-called signatures ‘‘L1’’ and ‘‘L2,’’ including the Ni-ligating cysteines, provided the basis for the currently accepted classification of [NiFe] hydrogenases (Vignais and Billoud 2007). In most cases, the L2 signature ends at a histidine residue (His536 in D. gigas), which marks the C-terminal endopeptidase cleavage site. In group 4 [NiFe] hydrogenases, represented by hydrogenase-3 of E. coli, and the related enzymes of Rhodospirillum rubrum, Methanosarcina barkeri, Methanobacterium thermoautotrophicum, and Pyrococcus furiosus, this histidine is replaced by an arginine residue

Gas channel

4

Glu321 [NiFe] Mg

Glu18 His536

Glu46

. Fig. 4.7 Model for the proton transfer pathway in Desulfovibrio gigas hydrogenase (PDB entry: 2FRV). The putative pathway extending from the [NiFe] site to the protein surface (at Glu46 and Glu321) is indicated by dashed lines. Selected water molecules are shown as red spheres. Cavities and the surface area of the protein are represented as dark gray meshes. See text for details

(reviewed by Tersteegen and Hedderich 1999). By X-ray crystallography, the C-terminal histidine has been identified as a ligand to an additional metal: a magnesium ion in periplasmic [NiFe] hydrogenases (Volbeda et al. 1995, 2012; Higuchi et al. 1997; Montet et al. 1997; Matias et al. 2001; Fritsch et al. 2011b) and an iron ion in the [NiFe(Se)] hydrogenase (Garcin et al. 1999; Marques et al. 2010). Dihydrogen and protons have to traverse a distance of 20–30 A˚ between the active site and the surface of the protein. Several proton pathways involving histidines, glutamates, carboxylate groups, and internal water molecules have been discussed (Fdez Galvan et al. 2008; Teixeira et al. 2008; Baltazar et al. 2012; Sumner and Voth 2012). However, a systematic investigation of these pathways is still outstanding. The only experimental information on proton transfer comes from a relatively small number of site-directed mutants (Dementin et al. 2004). One possible proton pathway, proposed for the D. gigas hydrogenase (> Fig. 4.7), implies two conserved glutamic acid residues (Glu18 and Glu46) in the large subunit and several water molecules located between the [NiFe] and the Mg site (Fontecilla-Camps et al. 2007). For a long time, the diffusion of molecular hydrogen, the smallest molecule in nature, through a protein matrix was considered to occur randomly. A cavity map, calculated from the electron density data for the crystallized D. gigas hydrogenase, showed a network of hydrophobic channels connecting the active site with the protein surface. Experiments on the diffusion of xenon atoms in crystals of the [NiFe] hydrogenase from D. fructosovorans indicated that there exist a few discrete hydrophobic cavities (> Fig. 4.5) which were interpreted as gas channels in the protein (Montet et al. 1997). More recent molecular simulation approaches revealed that the hydrophobic channel may account for only 60 % of the total H2

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4

H2-Metabolizing Prokaryotes

flux and that there is a diverse network of accessible sites, which may facilitate diffusion of H2 to the active site (Wang et al. 2011).

Catalytic Reaction Unlike platinum, hydrogenases operate by a heterolytic mechanism cleaving dihydrogen to H and H+ as deduced from early isotope exchange experiments (Krasna 1979). Although the precise reaction is still not fully understood, biochemical and X-ray structure analyses in concert with magnetic resonance techniques (EPR, ENDOR, ESEEM) and Fourier transform infrared (FTIR) spectroscopy, X-ray absorption spectroscopy (XAS), and density functional theory (DFT) calculations have shed light on the chemical mechanism of the catalytic cycle (reviewed by Fontecilla-Camps et al. 2007; Ogata et al. 2009). Spectroscopy of standard [NiFe] hydrogenases has shown that the unpaired electron, or the EPR-detectable unpaired spin, is located close to nickel and possibly in the vicinity of one of its sulfur ligands, but not on the iron. From this observation, it was inferred that the iron is retained as a low-spin Fe(II) during the entire catalytic cycle and that the diatomic ligands on the iron contribute to the maintenance of this redox state (Happe et al. 1997). The nickel at the active site undergoes several redox changes. Initially, three paramagnetic, EPR-detectable Ni-states, Niu-A, Nir-B, and Nia-C, were identified (> Fig. 4.8) which differed in the g values of the rhombic signals and in the infrared stretching frequencies. Aerobically isolated hydrogenase from D. gigas contains a mixture of the Niu-A and Nir-B forms which display different activation kinetics. The unready Niu-A state shows a Ni(III) signal with a gy value of 2.24 in the EPR spectrum.

Niu-A −96 mV

e−(H+)

This state can be fully activated only after incubation with H2 for several hours. In contrast, the ready Nir-B state, exhibiting a gy value of 2.16 in the Ni(III) signal, can be activated by H2 within a few minutes (Fernandez et al. 1985; Teixeira et al. 1985). These oxidized forms of the hydrogenase contain oxygen species as bridging ligands between the two transition metals (> Fig. 4.8), as shown by X-ray structure analysis (reviewed by Ogata et al. 2009). Both oxidized states are inactive and do not participate in the catalytic cycle. Sequential one-electron reductions by external reducing agents lead to EPR-silent Ni-S forms and finally result in the formation of the EPR-detectable catalytically active Nia-C state (van der Zwaan et al. 1990; > Fig. 4.8). Nia-C is photosensitive and can be converted to the EPR-detectable Nia-L state (Ni[I]) upon illumination at low temperatures. Binding of CO to the Nia-C state also yields a paramagnetic Ni-CO form (Ni[I]). Further reduction of Nia-C at low redox potentials leads to the EPR-silent Nia-SR states (> Fig. 4.8). During reductive activation of hydrogenase, the oxygen species dissociate most likely as water molecules from the [NiFe] active site, and the catalytic cycle is initiated. According to X-ray studies, the gas channel from the surface ends at the nickel, and the inhibitors CO and peroxide also bind at this metal (Ogata et al. 2002; Volbeda et al. 2005). Therefore, nickel has been suggested to be the initial site for H2 activation. This is still a controversial issue, since most DFTand model complex studies predict that H2 binds to the low-spin Fe(II) (Kubas 2007; Lill and Siegbahn 2009). The Ni-Fe bridging hydride that could be detected by advanced magnetic resonance spectroscopy (Brecht et al. 2003; Pandelia et al. 2012) is widely accepted as proof for the heterolytic cleavage of H2. One of the terminal cysteine ligands which is replaced by a selenocysteine in the [NiFe(Se)] hydrogenases is assumed to act as the primary base accepting the

Niu-A (unready)

Nir-B −151 mV

Niu-S Reduction

148

e−(H+)

Nir-S

+H+,−H2O

hn T < 100K

− + −375 mV e ,H

e−,H+

Nia-SR Nia-SR' Nia-SR"

CN CO

S

Nir-B (ready) OH−

S

CN Fe2+

Ni3+

Nia-C −436 mV

S

S

CN

Fe2+

Ni3+

active Nia-S

inactive Nia-L

O OH

S

S

S

CN CO

S

Nia-C (active) H−

S Ni3+ S

CN Fe2+

S

S

CN CO

. Fig. 4.8 Model for the activation and light sensitivity of standard [NiFe] hydrogenases. The nature of the bridging species in the Niu-A state is still a matter of debate. Apparent midpoint potentials (Em) of the respective redox transition are shown according to Fichtner et al. (2006). The structural models of selected [NiFe] oxidation states are depicted on the right. See text for details

H2-Metabolizing Prokaryotes

proton (> Fig. 4.7). To keep the catalytic cycle running, the fully reduced hydrogenase in the Nia-SR state (> Fig. 4.8) needs to be reconverted to a more oxidized H2-accepting species. Reoxidation is achieved by the release of protons and the transfer of electrons from the [NiFe] center via the Fe-S cluster(s) to an external redox partner (> Fig. 4.6). Once this reaction is completed, the enzyme is ready to enter the next catalytic cycle (Albracht 1994; Cammack 2001; Stein and Lubitz 2002). The scheme in > Fig. 4.8 does not apply to all [NiFe] hydrogenases, since several redox states are not detectable in representatives of non-standard hydrogenases (Saggu et al. 2009; Horch et al. 2010; Pandelia et al. 2010).

Interaction of [NiFe] Hydrogenases with Cytochromes The type of interaction with a specific cellular redox partner can usually be deduced from structural features of the hydrogenase small subunit. In the periplasmic hydrogenases of sulfate

4

reducers, the C-terminal histidine residue that is involved in coordination of the distal [4Fe-4S] cluster (> Fig. 4.6) is located close to the surface of the small subunit. Several species of Desulfovibrio contain periplasmic low-potential c-type cytochromes. Of these cytochromes, the tetraheme cytochrome c3 is considered as the primary redox partner of periplasmic hydrogenases (Matias et al. 2002; Yahata et al. 2006). The shuttling of electrons from the hydrogenase to membrane complexes of polyheme cytochromes of the Hmc family by c-type cytochromes may be a general mechanism in Desulfovibrio species (Rossi et al. 1993). Membrane-bound [NiFe] hydrogenases attached to the periplasmic side of the cytoplasmic membrane are found in many proteobacteria. They are characterized by a highly conserved (approx. 50 amino acid-long) oligopeptide at the C-terminus of the small subunit (> Fig. 4.9). This hydrophobic region is essential for binding the hydrogenase to the membrane and coupling the electron flow to the quinone pool of the respiratory chain. A membrane-integral cytochrome b has been isolated in complex with the [NiFe] hydrogenase from Wolinella

. Fig. 4.9 Modular structure and domain organization of selected [NiFe] hydrogenases. Gray boxes represent hydrogenase subunits (not drawn to scale). Redox partners are listed at the right. A key to the symbols is given in the lower part of the figure. (a) D. gigas periplasmic hydrogenase (Volbeda et al. 1995). (b) R. eutropha regulatory hydrogenase (Kleihues et al. 2000). The hatched area symbolizes the characteristic C-terminal region of the small subunit. (c) R. eutropha membrane-bound hydrogenase (Frielingsdorf et al. 2011; Fritsch et al. 2011b). The hatched area symbolizes the characteristic hydrophobic C-terminal region of the small subunit. (d) Methanosarcina barkeri Ech hydrogenase (Forzi et al. 2005). (e) Pyrococcus furiosus sulfhydrogenase (Silva et al. 1999). (f) R. eutropha soluble hydrogenase (Lauterbach et al. 2011a, b)

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H2-Metabolizing Prokaryotes

succinogenes (Dross et al. 1992, 1993), Ralstonia eutropha (Frielingsdorf et al. 2011), and Aquifex aeolicus (Guiral et al. 2005a). Interestingly, the high molecular mass complex isolated from R. eutropha appears to consist of three heterotrimeric units which may be relevant for intermolecular electron transfer (Frielingsdorf et al. 2011). The cytochrome from W. succinogenes, designated HydC, binds two heme groups, and analysis of mutants revealed that substitution of three histidine residues in HydC (His25, His67 and His186) predicted to be heme B ligands abolished quinone reactivity of the W. succinogenes hydrogenase, while benzylviologen reduction was retained. A similar phenotype was observed by mutating two conserved histidine residues in the small hydrogenase subunit HydA. One of the altered residues is located in the predicted membrane-integral C-terminal helix of HydA (His 305), and the other is likely involved in ligation of the distal [4Fe-4S] cluster. The data convincingly show that these components are necessary for electron transport from H2 to either fumarate or polysulfide and for quinone reactivity (Gross et al. 1998). Moreover, experiments with cytochrome b deficient mutants of R. eutropha demonstrated that the b-type cytochrome is bifunctional. In addition to its electron-transferring function, it anchors the hydrogenase to the membrane (Bernhard et al. 1997). A similar energy-converting system is operates in the archaeon Methanosarcina mazei Go¨ 1 (> Fig. 4.3). The outside-oriented membrane-bound [NiFe] hydrogenase of this strain transfers electrons from H2 oxidation via a b-type cytochrome and methanophenazine to heterodisulfide oxidoreductase. This reaction is coupled to the generation of a proton motive force (Ide et al. 1999). Thus, it appears that the cytochrome b serves as a common primary electron acceptor for a large group of hydrogenases.

A Histidine Kinase Is the Target for the HydrogenSensing [NiFe] Hydrogenases The H2-sensing regulatory [NiFe] hydrogenases of Bradyrhizobium japonicum (Black et al. 1994), Rhodobacter capsulatus (Elsen et al. 1996), and Ralstonia eutropha (Lenz et al. 1997) also contain a characteristic C-terminal region of approximately 50 amino acids in the small subunit. The sequence of this peptide is highly conserved within this group of soluble cytoplasmic proteins and completely distinct from the C-terminal region of the membrane-bound [NiFe] hydrogenase counterparts (> Fig. 4.9). This observation points to a specific role of the C-terminal extension in partner recognition and, in this case, in signal transduction. In fact, the formation of a tight complex between the H2-sensing hydrogenase of R. eutropha and its cognate signal-transmitting histidine protein kinase has been demonstrated in vitro using purified components (Bernhard et al. 2001). Unlike the standard energy-converting [NiFe] hydrogenases, which are isolated as simple heterodimers (ab), the H2-sensing hydrogenase of R. eutropha was obtained as a tetramer consisting of two heterodimeric species (a2b2).

A mutant deleted for the C-terminal peptide of the small subunit lost its H2-sensing ability but still catalyzed H2 oxidation. The mutant protein formed neither an a2b2 species nor a complex with the histidine protein kinase (Buhrke et al. 2004), supporting the notion that the C-terminus of the small subunit is essential for both complex formation and H2 signal transduction.

Multimeric [NiFe] Hydrogenases Less information is available on the interaction of the hydrogenase module within multisubunit [NiFe] hydrogenase complexes. These modules are often characterized by a truncated form of the small subunit, which differs markedly in its amino acid composition and cofactor content from the small subunit of the prototypic D. gigas hydrogenase (> Fig. 4.9). Heteromultimeric [NiFe] hydrogenases are generally soluble, residing in the cytoplasm, or are associated with the inner surface of the cytoplasmic membrane. A typical feature of one group of multimeric hydrogenases is a tight association of the hydrogenase module with a second redox-active moiety that binds coenzymes such as F420 (8-hydroxy-5-deazaflavin), NAD+, or NADP+, which are reversibly reduced by H2. The F420-reducing hydrogenases of methanogens are heterotrimeric FAD-containing enzymes that tend to form aggregates. A well-characterized example is the F420-reducing hydrogenase from Methanobacterium formicicum (Baron and Ferry 1989a, b), which consists of the subunits FrhA, FrhB, and FrhG. The FrhA subunit contains the [NiFe] center, and the FrhB subunit harbors the binding site for the cofactor F420. Variants of this hydrogenase exist which contain selenium in addition to nickel and iron (reviewed by Baltazar et al. 2011). Methylotrophic methanogens such as Methanosarcina strains possess membrane-bound multimeric hydrogenases (> Fig. 4.3). In Methanosarcina mazei, a cytochrome b serves as the primary electron acceptor (Ide et al. 1999). The redox carrier methanophenazine shuttles electrons from the hydrogenase to heterodisulfide reductase. In hydrogenotrophic methanogens, which are devoid of cytochromes, the F420nonreducing hydrogenase transfers electrons directly to an associated heterodisulfide reductase (Stojanowic et al. 2003). Methanococcus voltae harbors two enzymes of the [NiFe] type and two enzymes of the [NiFe(Se)] type. One enzyme of each pair is able to reduce F420, whereas the other species do not reduce the F420-coenzyme. The large subunit lacks the C-terminal cysteine, and instead, a selenocysteine provides the ligand to Ni. This binding site is contained on a separate peptide of 25 amino acids (Halboth and Klein 1992). The soluble cytoplasmic NAD-linked [NiFe] hydrogenases (reviewed by Burgdorf et al. 2005; Tamagnini et al. 2007; Horch et al. 2012) consist of up to six subunits (> Fig. 4.9). In the case of the soluble hydrogenase (SH) from R. eutropha, the hydrogenase module contains in addition to the active site subunit HoxH (52 kDa) a truncated form of the small, electrontransferring subunit HoxY (23 kDa) with only one Fe-S cluster

H2-Metabolizing Prokaryotes

as a prosthetic group and an additional loosely bound FMN (van der Linden et al. 2004). This module is associated with a heterodimeric iron flavoprotein, the so-called diaphorase (Schneider and Schlegel 1976; Schneider et al. 1984a; TranBetcke et al. 1990). The diaphorase consists of a large polypeptide, HoxF (67 kDa), and a small subunit, HoxU (26 kDa). The diaphorase moiety accommodates three to four iron-sulfur clusters and one FMN (Lauterbach et al. 2011a). Sequence alignments revealed a close relationship between the diaphorase part of the NAD-linked hydrogenases and three peripheral subunits of bacterial and mitochondrial complex I (Tran-Betcke et al. 1990; Pilkington et al. 1991; Friedrich et al. 2000). The HoxF polypeptide appears to be a fusion product of the NuoE and NuoF subunits of complex I, and HoxU is homologous to the Nterminal part of NuoG (reviewed by Vignais and Billoud 2007; Efremov and Sazanov 2012). In addition, the SH comprises a dimer of HoxI (19 kDa) subunits associated with the diaphorase. HoxI contains a putative nucleotide binding site, similar to that of cAMP receptors, and is assumed to act as an alternative electron entry site for reductive reactivation of the SH via NADPH (Burgdorf et al. 2005). The NAD-reactive hydrogenases are found in aerobic H2-oxidizing bacteria including the well-studied Rhodococcus opacus (formerly Nocardia opaca) and Ralstonia eutropha (formerly Alcaligenes eutrophus; Schneider and Schlegel 1976; Schneider et al. 1984a; Tran-Betcke et al. 1990; Grzeszik et al. 1997a). A cytoplasmic, NAD-reducing hydrogenase was identified in the methanotroph Methylococcus capsulatus (Hanczar et al. 2002). Similar enzymes, designated ‘‘bidirectional hydrogenases,’’ have also been found in cyanobacteria (reviewed by Tamagnini et al. 2007). An NAD-reactive cyanobacterial hydrogenase from the filamentous Anabaena variabilis ATCC 29413 was the first enzyme of this type to be characterized and sequenced (Schmitz et al. 1995; Serebryakova et al. 1996). Subsequently, this type of hydrogenase was also identified in unicellular cyanobacteria and appears to be loosely associated with cytoplasmic and thylakoid membranes (Kentemich et al. 1989; Serebryakova et al. 1994). The bidirectional hydrogenase of Synechocystis sp. PCC 6803 was proposed to serve as an electron valve during photosynthesis (Appel et al. 2000) and later shown to have a bias toward proton reduction (McIntosh et al. 2011). Unlike the SH from R. eutropha, the diaphorase moiety of these cyanobacterial-type hydrogenases is associated with an additional subunit designated ‘‘HoxE’’ (Schmitz et al. 2002) that is homologous to NuoE, the [2Fe-2S]-containing subunit of complex I. HoxE could be involved in membrane interactions or electronic coupling between the hydrogenase and diaphorase moiety (Tamagnini et al. 2007). Two bidirectional hydrogenases, Hox1 and Hox2, were discovered in the purple sulfur bacterium Thiocapsa roseopersicina. Hox1 appears to be connected to sulfur metabolism and fermentative processes in the dark, and Hox2 evolves hydrogen in vivo in the presence of glucose at low sodium thiosulfate levels (Ra´khely et al. 2007; Maro´ti et al. 2010). A group of closely related tetrameric NADP-reactive [NiFe] hydrogenases has been found in the hyperthermophilic

4

archaeon Pyrococcus furiosus (Ma et al. 1993, 2000; Pedroni et al. 1995) and Thermococcus litoralis (Ra´khely et al. 1999). The HydA and HydD subunits of this type of enzyme constitute the hydrogenase module, whereas the HydB and HydG subunits form the flavin-containing NADP-reactive moiety of the protein (> Fig. 4.9). Early studies suggest that this group of [NiFe] hydrogenases has both H2-oxidizing and S0-reducing activities (Ma et al. 1993). Depending on the pH, S0 reduction activity of the purified hydrogenase from P. furiosus is increased by addition of the redox protein rubredoxin to the assay (Ma et al. 1993). The authors propose that during fermentation, the hydrogenase can dispose of excess reductant using either protons or S0/polysulfides as the electron acceptor and therefore designated it a ‘‘sulfhydrogenase.’’ Alternatively, a putative soluble NADPH-sulfur oxidoreductase may be responsible for the S0 reduction activity in P. furiosus (Jenney and Adams 2008), and recent studies indicate that a membrane-bound [NiFe] hydrogenase is accountable for the major H2 production under sulfurlimited conditions (Schut et al. 2012). The reduction of elemental sulfur with electrons derived from molecular hydrogen drives chemolithoautotrophic growth of the hyperthermophilic and acidophilic archaeon Acidianus ambivalens under anaerobic conditions. The sulfur respiration is mediated by a membrane-bound [NiFe] hydrogenase that transfers the electrons from H2 oxidation via the quinone pool to sulfur reductase (Laska et al. 2003). The hyperthermophilic bacterium Aquifex aeolicus can also grow on H2 and S0, producing H2S as the metabolic end product. In A. aeolicus, the electron transfer is mediated by a membrane-bound supercomplex that was enriched and shown to contain two different membrane-bound periplasmic [NiFe] hydrogenases, a sulfur reductase from the molybdoprotein family and components of the cytochrome bc1 complex (Guiral et al. 2005b).

Energy-Converting [NiFe] Hydrogenases A special subfamily of the multimeric [NiFe] hydrogenases has been designated ‘‘energy-converting’’ hydrogenases (reviewed by Hedderich and Forzi 2005; Thauer et al. 2010). In addition to the large and small hydrogenase subunits, they usually contain two hydrophilic and two membrane-integral subunits. These five to six subunits form the basic membrane-bound core of the energy-converting hydrogenases that is conserved in all members of this group. The core shares sequence similarity with the NuoBCDIHL subunits, the catalytic core of complex I (Efremov and Sazanov 2012). Several energyconverting [NiFe] hydrogenase complexes are capable of coupling the oxidation of carbonyl groups from formate, acetate, or carbon monoxide to the reduction of protons. This exergonic reaction is linked to energy conservation by means of electrontransport phosphorylation (Hedderich and Forzi 2005). Other representatives of this group couple the endergonic reduction of low-potential ferredoxins using H2 as electron donor in a reaction driven by reverse electron transport (Thauer et al. 2010).

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A well-studied example of the energy-converting [NiFe] hydrogenases is the formate hydrogenlyase (FHL) complex in E. coli that catalyzes the conversion of formate into CO2 and H2 (Bo¨hm et al. 1990; Sauter et al. 1992). FHL is anticipated to comprise seven hyc gene products (Forzi and Sawers 2007). The hydrogenase module of FHL, the so-called hydrogenase-3 (Hyd-3), consists of the active site containing subunit HycE and the small subunit HycG, which harbors only the proximal [4Fe-4S] cluster (> Fig. 4.2). Much of which is known today about the maturation of the [NiFe] center in hydrogenases has been deduced from experiments with Hyd-3 of E. coli (see below). Both the formate dehydrogenase as well as the two hydrogenase subunits are attached to the inner side of the cytoplasmic membrane via membrane proteins, which serve as membrane anchors and electron mediators between the two redox proteins (Bo¨hm et al. 1990; Sauter et al. 1992). Due to the sequence similarity with complex I, a possible role of FHL in energy conservation under fermentative conditions has been proposed (Sauter et al. 1992). However, so far no clear experimental evidence is available. Due to its high instability, biochemical data on the complete FHL complex from E. coli is still limited. A similar type of six subunit FHL core was successfully purified recently from the hot spring bacterium Klebsiella oxytoca (Wu et al. 2011). The isolated protein complex exhibited high H2 production activity, and mutant studies suggest that this hydrogenase is the main enzyme responsible for hydrogen production in K. oxytoca under O2 stress conditions. The CO-induced [NiFe] hydrogenase of the photosynthetic bacterium Rhodospirillum rubrum is a constituent of a large enzyme complex that couples the oxidation of CO to the reduction of protons permitting the organism to grow in the dark with CO as the sole energy source (Uffen 1976). Electron transfer between the CO dehydrogenase (CooS) homodimer and [NiFe] hydrogenase (CooHL) is mediated by the cytoplasmic 2[4Fe-4S] ferredoxin CooF (Singer et al. 2006). Like E. coli FHL, the CO oxidizing/H2 evolving complex from R. rubrum is rather labile (Fox et al. 1996a, b). Nevertheless, an intact and highly active CooFHKLMUSX complex could be isolated from Carboxydothermus hydrogenoformans (Soboh et al. 2002; > Fig. 4.2). It has been proposed that the CO-induced hydrogenase couples CO-dependent H2 evolution to proton translocation to produce a proton gradient (Fox et al. 1996b). CO-dependent ATP synthesis has been demonstrated in vivo in Rubrivivax gelatinosa (Maness et al. 2005). This photosynthetic bacterium possesses a CO-dependent H2 evolution system that is very similar to that in R. rubrum. Energy-converting [NiFe] hydrogenases in some methanogens, designated Ech hydrogenase, contain in addition to the six conserved core subunits up to 14 accessory subunits (Thauer et al. 2010). An intact EchABCDEF hydrogenase complex (> Fig. 4.9) has been purified to homogeneity from Methanosarcina barkeri cells (Meuer et al. 1999). It is composed of two membrane-integral (EchAB) and four hydrophilic subunits (EchCDEF) and catalyzes the reversible reduction of a lowpotential 2[4Fe-4S] ferredoxin with H2 (Meuer et al. 1999). With

the aid of a mutant, it was elegantly shown that Ech hydrogenase has a key function in methanogenesis in M. barkeri (Meuer et al. 2002). When the cells grow on acetate, Ech is supposed to mediate H2 evolution from ferredoxin that is reduced by the oxidation of an acetate-borne CO group to CO2 (Hedderich and Forzi 2005). During autotrophic growth on CO2 and H2, Ech catalyzes the energetically unfavorable reduction of ferredoxin, which in turn is used as a low-potential electron donor for various soluble oxidoreductases, for example, the acetyl-CoA synthase/CO dehydrogenase complex, pyruvate-ferredoxin oxidoreductase, and formylmethanofuran dehydrogenase (Hedderich and Forzi 2005). The reduction of CO2 to formylmethanofuran is the first step of methanogenesis. Because of the low midpoint potential of the CO2 + methanofuran/ formylmethanofuran couple (E00 = 500 mV), the reaction is endergonic even with H2 as the electron donor (E00 = 414 mV; Bertram and Thauer 1994). Based on biochemical and genetic studies, a function for Ech hydrogenase as proton or sodium pump has been proposed (Bott and Thauer 1987; Kaesler and Scho¨nheit 1989; Meuer et al. 1999; Albracht and Hedderich 2000). The observed pH dependence of EPR signals assigned to two [4Fe-4S] clusters in the EchC and EchF subunits of Ech hydrogenase suggests that the two iron-sulfur clusters actively participate in the proton-translocating process (Forzi et al. 2005). Using inverted membrane vesicles, the ferredoxindependent Ech hydrogenase from Pyrococcus furiosus was shown to couple H2 evolution and ATP synthesis (Sapra et al. 2003).

[FeFe] Hydrogenases Molecular Characteristics The [FeFe] hydrogenases (formerly known as ‘‘Fe-only hydrogenases’’) are found in anaerobic, primarily H2-evolving organisms, including fermentative bacteria, sulfate-reducers, and some lower eukaryotes. They are characterized by extremely high oxygen sensitivity, a high turnover rate, and a low affinity for the substrate hydrogen (reviewed by Fontecilla-Camps et al. 2007, Mulder et al. 2011 and Vignais and Billoud 2007). In addition to monomeric [FeFe] hydrogenases, heterodimeric, -trimeric and -tetrameric enzymes exist. Two well-characterized examples are the monomeric 61-kDa cytoplasmic [FeFe] hydrogenase of Clostridium pasteurianum (Adams and Stiefel 1998) and the heterodimeric periplasmic enzyme of Desulfovibrio desulfuricans consisting of a large (42kDa) and a small (11-kDa) subunit (Nicolet et al. 1999). The [FeFe] hydrogenase of C. pasteurianum uses protons as electron acceptor to dispose of excess reducing equivalents, thereby regenerating oxidized ferredoxin and producing H2. The periplasmic [FeFe] hydrogenase of D. desulfuricans plays a role in H2 uptake. The [FeFe]-containing active site consists of a unique metal-containing prosthetic group, the so-called H-cluster. Multiple amino acid sequence alignments revealed considerable similarity within the H-cluster-coordinating region, indicating

H2-Metabolizing Prokaryotes

a

2

DdH

397 421 1

F

L

1

CpI

123

S 511 516

Fd

35

4

517 522

574

F H-domain

b

N terminus small subunit C terminus large subunit

N terminus

C terminus small subunit C terminus N terminus large subunit

Non-protein cysteine

DdH

CpI

. Fig. 4.10 Structural comparison of two [FeFe] hydrogenases. (a) Schematic representation of the various domains of the Desulfovibrio desulfuricans hydrogenase (DdH) and the Clostridium pasteurianum hydrogenase I (CpI). Structurally related domains are depicted in the same color. The small and large subunits of DdH are labeled S and L, respectively. The homologous parts of the H-domain are colored dark blue and red. The dotted yellow line indicates a gap in the sequence alignment arising from peptide insertions at the N- and Cterminal ends of the small and large subunits of the DdH, respectively. The [2Fe-2S] plant-ferredoxin-like (Fd) domain is shown in green and the 2[4Fe-4S] ferredoxin-like F-domain (F) in turquoise. An atypical [4Fe-4S]-cluster-containing domain is colored pink. (b) The threedimensional structures of DdH and CpI with the same colors as in (a) (Adapted from Nicolet et al. (2000))

a fairly conserved architecture of the active site. Additional domains, however, which differ markedly in size and cofactor content, are often present in this class of hydrogenases (reviewed by Vignais and Billoud 2007). The three-dimensional structures of hydrogenase isoenzyme I from C. pasteurianum (CpI) (Peters et al. 1998) and the [FeFe] hydrogenase from D. desulfuricans (DdH) (Nicolet et al. 1999) have been solved at a resolution of 1.8 and 1.6 A˚, respectively. Comparison revealed a highly related multi-domain structure divided into an active site domain (H-domain), bearing the Hcluster, and accessory cluster domains (> Fig. 4.10). Both hydrogenases share in addition to the H-domain the so-called F-domain, a ferredoxin-like domain containing two [4Fe-4S] centers (‘‘F-clusters’’) (Peters et al. 1998; Nicolet et al. 2000). In the DdH, both the large and the small subunits form the H-domain and are topologically equivalent to the single chain of CpI. As a consequence, the C-terminus of the large and the N-terminus of the small DdH subunit are very close in space. The equivalent regions in CpI are connected by a loop (> Fig. 4.10b, yellow). The CpI contains two additional Fe-S centers that are absent in the DdH. Both centers are coordinated at N-terminal domains of the protein (> Fig. 4.10, green and

pink). One of these is a [2Fe-2S] cluster-containing domain with striking similarity to plant-type ferredoxins (Nicolet et al. 2000). This plant-type ferredoxin is connected to the F-domain via a rather short region that coordinates a [4Fe-4S] cluster through three cysteine ligands and a single histidine (Peters et al. 1998). This coordination resembles the distal [4Fe-4S] cluster coordination in periplasmic [NiFe] hydrogenases where the histidine ligand plays a crucial role in the intra- and intermolecular electron transfer properties of the protein (Dementin et al. 2006).

Catalytic Reaction The H-cluster is composed of a typical [4Fe-4S] cubane that is connected to a 2Fe subcluster by a single cysteine residue (Nicolet et al. 1999; Peters et al. 1998). An outstanding feature of the 2Fe subcluster of the H-cluster discovered in both the C. pasteurianum and the D. desulfuricans [FeFe] hydrogenases is the existence of five diatomic ligands and a nonprotein fiveatom dithiolate ligand bridging Fe1 and Fe2 (> Figs. 4.4, > 4.11). Early FTIR studies of [FeFe] hydrogenases suggested

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H2-Metabolizing Prokaryotes

Hox-CO HN

OC

S S

Hred

Hox HN

HN

S

Fe22+ Fe11+ Cys C OC CO CN NC O

[4Fe-4S]2+

[4Fe-4S]2+

[4Fe-4S]2+ +CO

S S

H2O Fe2 OC

2+

NC

C O

+

+H2,–H

S 1+

Fe1

CN

S S

H

Cys

1+

Fe2 OC

CO

NC

C O

S Fe11+ Cys CN

CO

. Fig. 4.11 Structural models of selected H-cluster oxidation states in bacterial [FeFe] hydrogenase. The models are based on crystal structures of the C. pasteurianum hydrogenase (Peters et al. 1998; Lemon and Peters 1999) and advanced magnetic resonance spectroscopy as well as theoretical investigations (Siegbahn et al. 2007; Silakov et al. 2007, 2009). See text for details

the presence of both CN and CO ligands (Pierik et al. 1998) thus providing a similar coordination environment as found in [NiFe] hydrogenases. Unlike the [NiFe] center, which is bound to the protein by four cysteine ligands, only Fe1 of the 2Fe subcluster is covalently linked to the protein via a single cysteine (> Fig. 4.11). The two Fe atoms are bridged through a CO ligand and the five-atom dithiolate ligand that was tentatively identified as a propanedithiol and later as a dithiomethylether in the X-ray studies (Nicolet et al. 1999; Pandey et al. 2008). Recent spectroscopic and theoretical investigations provided strong evidence for a di-(thiomethyl)amine group (–S–CH2–NH– CH2–S–) as the bridging ligand (Ryde et al. 2010; Silakov et al. 2009). A terminal water molecule was identified as a fifth ligand at Fe2 in the C. pasteurianum hydrogenase structure (Peters et al. 1998; Pandey et al. 2008). As postulated for the [NiFe] site, the strong field ligands CO and CN are anticipated to stabilize the two iron ions at low oxidation states. Consequently, the 2Fe subcluster of oxidized [FeFe] hydrogenases is in a low-spin S = ½ state with an Fe(1+)/Fe(2+) pair (Silakov et al. 2007, > Fig. 4.11). The coordination environment of the two metals favors the distal Fe2 as a candidate for displacement and formation of a bound hydride intermediate. This assumption is consistent with the observation that the competitive inhibitor CO binds reversibly to Fe2 in CpI (Lemon and Peters 1999; > Fig. 4.11). Further support comes from the observation that the putative gas diffusion pathways end at the Fe2 coordination site of the CpI and DdH structures (Cohen et al. 2005a, b). This region appears to be highly conserved in [FeFe] hydrogenases pointing to a specific access of H2 to the active site as has been discussed for the [NiFe] hydrogenases (Montet et al. 1997). The exact catalytic mechanism of H2 conversion at the H-cluster is still discussed controversially. According to the current understanding (reviewed by Fontecilla-Camps et al. 2007; Lubitz et al. 2007; Mulder et al. 2011), the proximal Fe1 of the H-cluster remains in the formal +1 state in Hox and Hred (> Fig. 4.11), whereas only the distal Fe2 alternates between the formal +2 (ox) and +1 (red) state. This model is assumed to account for H2 oxidation as well as H+ reduction. X-ray and FTIR studies indicate that the Fe-Fe bridging CO ligand becomes terminal when switching from Hox to Hred (> Fig. 4.11). After heterolytic cleavage of H2 at Fe2 (during H2 uptake), the hydride

Glu282 Ser319

Cys299 H2O Glu279 Fe1

Fe2

H-Cluster

. Fig. 4.12 Putative proton transfer pathway from the [FeFe] active site of C. pasteurianum hydrogenase (PDB entry: 3C8Y) to the protein surface (Adapted from Cornish et al. (2011)). The proton path is indicated by black dashed lines. The surface area of the protein is represented as dark gray meshes. See text for details

intermediate remains at Fe2. The released proton is initially attached to a nearby base, likely the di-(thiomethyl)amine group. A potential proton transport pathway in the C. pasteurianum enzyme involves a water molecule and four amino acid residues (Cys299, Glu279, Ser319, and Glu282) that constitute the connection of the H-cluster to the protein surface (> Fig. 4.12). These residues are strictly conserved in all [FeFe] hydrogenases, and their importance in catalysis was recently demonstrated through substitutions via site-directed mutagenesis resulting in a drastic reduction of hydrogenase activity (Cornish et al. 2011). It is likely that electron transfer from and/or to the H-cluster is regulated via the accessory F-clusters (Peters et al. 1998; > Fig. 4.10).

Diversity of [FeFe] Hydrogenases and Related Proteins The smallest [FeFe] hydrogenase unit (45–48 kDa) consisting only of an H-cluster domain has been discovered in green algae (Happe et al. 1994; Happe and Naber 1993). This type of enzyme, first reported for Chlamydomonas reinhardtii

H2-Metabolizing Prokaryotes

4

. Fig. 4.13 Modular structure and domain organization of selected [FeFe] hydrogenases (Adapted from Vignais and Billoud (2007)). Gray boxes represent hydrogenase subunits (not drawn to scale). Redox partners are listed on the right. A key to the symbols is given at the bottom of the figure

(> Fig. 4.13), appears to be present also in Scenedesmus obliquus (Florin et al. 2001; Wunschiers et al. 2001) and Chlorella fusca (Winkler et al. 2002). The hydrogenase receives the electrons for H2 evolution from a [2Fe-2S] ferredoxin, which is reduced during the fermentative metabolic cycle of these organisms (Florin et al. 2001). Remarkably, sulfur-deprived cultures of C. reinhardtii were shown to perform light-dependent H2 production, indicating that under these conditions, [FeFe] hydrogenase can be linked to the photosynthetic electron-transport chain (Melis et al. 2000). Another relatively simple [FeFe] hydrogenase, which contains in addition to the H-cluster a 2 [4Fe-4S] ferredoxin-like F-domain, is found in prokaryotic species such as Megasphera elsdenii (Atta and Meyer 2000; > Fig. 4.13) as well as in lower eukaryotes such as Trichomonas vaginalis (Bui and Johnson 1996; Horner et al. 2000). In spite of the absence of additional prosthetic groups, the M. elsdenii hydrogenase uses electron donors of the type found in clostridia including a 2[4Fe-4S] ferredoxin and a flavodoxin (Atta and Meyer 2000). In the anaerobic eukaryotic organisms, the [FeFe] hydrogenases are localized in intracellular organelles of endosymbiotic origin (Mu¨ller 1993; Martin and Muller 1998; Moreira and Lopez-Garcia 1998). In the case of chytrid fungi,

anaerobic ciliates, and trichomonads, these organelles are known as ‘‘hydrogenosomes.’’ They are considered as modified mitochondria that have lost the capacity for oxidative phosphorylation while gaining the ability of ferredoxin-coupled H2 production from pyruvate. The periplasmic uptake [FeFe] hydrogenase of Desulfovibrio vulgaris Hildenborough (> Fig. 4.13) that shares high sequence identity with the [FeFe] hydrogenase from D. desulfuricans consists of a small subunit (HydB, 13.5 kDa) bearing a twin-arginine signal peptide at the N-terminus and a large subunit (HydA, 46 kDa). Both subunits undergo processing during membrane translocation (Hatchikian et al. 1999). The D. vulgaris [FeFe] hydrogenase is upregulated in response to oxidative stress leading to the interpretation that this hydrogenase has a protective role (Fournier et al. 2004). Multisubunit [FeFe] hydrogenases were identified in several prokaryotic species (Vignais and Billoud 2007). The heterotrimeric [FeFe] hydrogenase from the hyperthermophilic, anaerobic bacterium Thermotoga maritima (> Figs. 4.2, > 4.13) is an example of the new class of bifurcating hydrogenases (reviewed by Buckel and Thauer 2012). During fermentation of carbohydrates, the enzyme oxidizes ferredoxin and NADH

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H2-Metabolizing Prokaryotes

synergistically at an about 1:1 ratio and produces H2 (Schut and Adams 2009). Coupling to the exergonic oxidation of ferredoxin (E00 = 453 mV) drives the thermodynamically unfavorable oxidation of NADH (E00 = 320 mV) to produce H2 (E00 = 423 mV). As a consequence, oxidized coenzymes are regenerated. On the basis of sequence analysis and EPR spectroscopic data (Verhagen et al. 1999), at least ten prosthetic groups including the H-cluster were assigned to the three hydrogenase subunits (> Fig. 4.13). Each subunit contains a domain that is similar to the [2Fe-2S] NuoE-like polypeptide of complex I. An additional domain, homologous to the NuoF subunit of complex I, was identified in the amino acid sequence of HydB (> Fig. 4.13). These observations emphasize the evolutionary relationship between complex I and certain [FeFe] hydrogenases (reviewed by Vignais and Billoud 2007). The heterotetrameric cytoplasmic NADP-reducing [FeFe] hydrogenase of D. fructosovorans has an overall structure that resembles the counterpart of Thermotoga maritima showing the typical complex I-related domains (> Fig. 4.13). The catalytic subunit is very similar to the monomeric clostridial hydrogenase (Malki et al. 1995), and the thioredoxin-like fold of the C-terminal domain of the NuoE-related HndA subunit has been confirmed by a nuclear magnetic resonance (NMR) solution structure (Nouailler et al. 2006). It has been suggested that the HndB and HndC subunits of D. fructosovarans are fused to a single polypeptide (HydB) in T. maritima (Vignais et al. 2001). Sequences related to the [FeFe] hydrogenase catalytic subunit have been identified in genomes of aerobic eukaryotes,

including the human genome. Members of this class of nuclear proteins are termed ‘‘nuclear prelamin A recognition factors’’ (NARFs; Barton and Worman 1999). Since H2-converting processes have so far not been observed in higher eukaryotes, it is unlikely that NARF proteins are implicated in energy metabolism. A NARF protein in Saccharomyces cerevisiae (Nar1p) was shown to be essentially required for maturation of cytosolic and nuclear Fe-S proteins (Balk et al. 2004).

[Fe] Hydrogenases Molecular Characteristics The [Fe] hydrogenases (formerly known as ‘‘metal-free’’ or ‘‘FeS-cluster-free’’ hydrogenases) of methanogenic archaea represent the third, phylogenetically distinct class of hydrogenases (reviewed by Thauer et al. 2010). These homodimeric enzymes, characterized as H2-forming methylenetetrahydromethanopterin dehydrogenase (Hmd), are present in methanogenic archaea in which they catalyze a step in methanogenesis. The catalytic reaction involves the reversible reduction of methenyltetrahydromethanopterin (methenyl-H4MPT+) to methyleneH4MPT and a proton by transferring a hydride ion to the proR side of the C14a carbon of methenyl-H4MPT (> Fig. 4.14b; Thauer 1998). The reduced product is used by the F420-dependent methylene-H4MPT dehydrogenase for the reduction of F420 (> Fig. 4.3). Both enzymes are coordinately

c a

Cys O

S OC

Fe2+

N O

C O HO

GMP

*Fe-GP

b

# R

O HN H2N

N

H

14a

• N H + N CH3 H N H

CH3 H

Methenyl-H4MPT+

+ H2

*

pro-R R pro-S H 14a H O • N H N CH3 + H+ HN H H2N

N

N H

CH3 H

#Methylene-H MPT 4

. Fig. 4.14 Function and crystal structure of the [Fe] hydrogenase from M. jannaschii. (a) Structure of the iron-guanylylpyridinol cofactor (Fe-GP). (b) [Fe] hydrogenases catalyze the reversible transfer of a hydride from H2 into the pro-R side of methenyl-H4MPT+ yielding methyleneH4MPT and a proton. The methenyl C14a has carbocation character and serves as hydride acceptor. (c) Structure of the [Fe] hydrogenase-substrate complex (according to Hiromoto et al. 2009a [PDB entry: 3F47]; Hiromoto et al. 2009b [PDB entry: 3H65]). The cofactors are shown in ball-and-stick representation

H2-Metabolizing Prokaryotes

expressed at a high level under nickel depletion (Afting et al. 1998). With the aid of hydrogen isotope assays and twodimensional NMR, it was possible to explore the H2-forming reaction in great detail including the stereoselective hydride transfer (Thauer et al. 1996). Unlike other hydrogenases, which are Fe-S proteins, the Hmd monomers contain only one redox-active iron bound to a thermolabile and light sensitive cofactor. Exposure of the Hmd’s to UV-A or blue light results in inactivation of the enzyme and release of iron (Buurman et al. 2000; Lyon et al. 2004b). The crystal structure of the Hmd from Methanocaldococcus jannaschii (Shima et al. 2008) finally revealed the nature of the labile active site cofactor (> Fig. 4.14a). The mononuclear iron is coordinated by a cysteine residue, two CO molecules, and the nitrogen of a guanylylpyridinol compound with back-bonding properties similar to those of cyanide. The structural arrangement of these ligands is similar to the low-spin iron in [NiFe] and [FeFe] hydrogenases. Thus, hydrogenases are an impressive example of convergent evolutionary development. The fifth ligand to the Fe in Hmd from M. jannaschii was initially obscure (Shima et al. 2008). Later, the crystal structure of an Hmd mutant variant and mass spectrometry as well as infrared spectroscopy provided evidence for a biologically unique acylcarbon-iron ligation at this coordination site (> Fig. 4.14a; Hiromoto et al. 2009a; Shima et al. 2012).

Catalytic Reaction Little is known about the catalytic mechanism of H2 conversion at the iron-guanylylpyridinol (Fe-GP) cofactor. Mo¨ssbauer spectroscopy of native, CO- or CN -inhibited Hmd and of the isolated Fe-GP cofactor indicates that the protein contains low-spin Fe(2+) that is not redox active (Shima et al. 2005). Nevertheless, inhibition of Hmd by CO suggests that the iron is involved in H2 activation (Lyon et al. 2004a). The Hmd catalytic reaction relies on a ternary complex (Lyon et al. 2004a). Therefore, this type of hydrogenase does not catalyze the three reactions characteristic for the other two classes of hydrogenases: (1) H2-dependent reduction of dyes, (2) the exchange of H+ from H2O into H2, and (3) ortho-para H2 conversion. However, in the presence of the hydride-accepting methenyl-H4MPT+, Hmd hydrogenases catalyze reactions (2) and (3) with kinetics similar to those of the [NiFe] and [FeFe] isoenzymes (Vogt et al. 2008). A catalytic model consistent with these results was deduced from the crystal structure of the [Fe]-hydrogenasemethylene-H4MPT complex (> Fig. 4.14c; Hiromoto et al. 2009b).

H2 Conversion in the Presence of O2 Despite the fact that methanogenesis occurs in anoxic environments, the [Fe] hydrogenases remain catalytically active in the presence of O2 (Lyon et al. 2004a). This behavior is in marked contrast to the highly O2-sensitive [FeFe] hydrogenases that

4

inactivate irreversibly by even trace amounts of O2 (De Lacey et al. 2007; Goldet et al. 2009; Stripp et al. 2009). The less O2-sensitive [NiFe] hydrogenases react in most cases reversibly with O2 giving rise to a mixture of the inactive ‘‘dormant’’ states Niu-A and Nir-B and related EPR-silent species (> Fig. 4.8). Enzymes in the Nir-B state reactivate rapidly under mildly reducing conditions, whereas the Niu-A state requires long-term reactivation that may occur exclusively in vitro (De Lacey et al. 2007; Vincent et al. 2007). Consequently, the strict avoidance of Niu-A-related states and the continuous removal of oxygen species related to Nir-B are prerequisites for a [NiFe] hydrogenase to function in vivo under aerobic conditions (Cracknell et al. 2009; Ludwig et al. 2009). Oxygen tolerance is a characteristic feature of a group of [NiFe] hydrogenases present in knallgas bacteria, which derive energy from oxidation of H2 in the presence of O2. Ralstonia eutropha H16 is a well-studied model organism of this group. The b-proteobacterium harbors at least three distinct [NiFe] hydrogenases capable of converting H2 even at ambient O2 levels (reviewed by Lenz et al. 2010). Oxygen tolerance is of major relevance for biotechnological application of hydrogenases as catalysts in various processes (Friedrich et al. 2011). Recent investigations focused on the elucidation of the molecular mechanism of oxygen tolerance. A well-characterized example of an O2-tolerant enzyme is the heterotrimeric membrane-bound [NiFe] hydrogenase (MBH) which is attached to the periplasmic side of the cytoplasmic membrane feeding electrons from H2 oxidation via a membrane-integral b-type cytochrome to the quinone pool of the respiratory chain (> Fig. 4.2). Comprehensive biochemical, spectroscopic, protein film electrochemistry and X-ray crystallography analyses on the MBH (Fritsch et al. 2011b; Goris et al. 2011) and its counterpart from Escherichia coli (Lukey et al. 2011; Volbeda et al. 2012) and Aquifex aeolicus (Pandelia et al. 2011) demonstrated that in these enzymes, the proximal Fe-S center differs in its electronic and molecular structure from the conventional [4Fe-4S] cubanes usually found at the corresponding position of O2-sensitive standard [NiFe] hydrogenases. The proximal [4Fe-3S] cluster of the MBH (> Fig. 4.15) is coordinated by two additional cysteine residues that are crucial for O2-tolerance of the protein. The unique architecture of this newly discovered cofactor enables it to undergo two redox transitions within an extraordinary narrow potential window. This allows the transfer of two instead of only one electron under physiologically relevant conditions (Goris et al. 2011). The redox changes are accompanied by significant structural rearrangements of the [4Fe-3S] cluster (Shomura et al. 2011). Therefore, the extended redox capability of the Fe-S cluster relay rather than a modified [NiFe] active site provides the key to understanding O2 tolerance of the MBH. According to the model, O2 is efficiently neutralized at the active site by rapid reduction to water, thereby preventing inactivation of the enzyme (> Fig. 4.15). Two crystal structure analyses of MBH proteins from knallgas bacteria (Fritsch et al. 2011b; Shomura et al. 2011) suggest a controlled translocation of water molecules from the

157

158

4

H2-Metabolizing Prokaryotes

[NiFe]

Em: −110 mVa

2e−, 2H+

Rapid Reactivation

e−, H+ H2O

[4Fe-3S]

Nir-B CN

S

Active Enzyme

Fe2+

Ni2+ S

CN

OH

S

Em: +160mVa/-60mVa

CO

S CN

S

Nia-S

H2

CN Fe2+

Ni3+

CO

S

S

S

−

[3Fe-4S]

H2O O2

Em: +25mVa

3H+, 3e−

[4Fe-4S]

Em: −180 mVa Cytochrome b Em: +10/+166 mVb Quinone pool Em: +90 mVb

. Fig. 4.15 A simplified model of the reactions of MBH with H2 and O2 (Adapted from Cracknell et al. (2009)). When attacking the MBH active site during H2-catalysis, O2 is immediately reduced with four electrons provided by a modified electron transfer relay in MBH, thereby producing one H2O and a [NiFe]-bridging hydroxide (Nir-B). After rapid reactivation with an additional electron, the enzyme reenters the catalytic cycle. The apparent midpoint potentials (Em) of the R. eutropha MBH cofactors that control the electron flow are taken from Knu¨ttel et al. (1994) (a) and Bernhard et al. (1997) (b)

active site to the protein surface via an extended network of hydrophilic cavities (> Fig. 4.16), supporting the notion that O2-tolerant hydrogenases have also an oxidase activity and form water as a byproduct during H2 conversion in the presence of O2. Similar conclusions were drawn from the crystal structure analyses of hydrogenase-1 from E. coli (Volbeda et al. 2012). In the case of the cytoplasmatic soluble hydrogenase (SH) from R. eutropha, a modified [NiFe] active site carrying additional cyanide ligands was previously suggested (Bleijlevens et al. 2004; van der Linden et al. 2004) to play a crucial role in the O2 tolerance of this enzyme. However, this was later proven wrong on the basis of in situ FTIR spectroscopy studies of the SH (Horch et al. 2010). The old observation that the isolated SH can be rapidly reactivated by traces of NADH (Schneider and Schlegel 1976) provides a more reasonable explanation for its O2 tolerance. Similar to the mechanism of the MBH, reverse electron transfer to the [NiFe] active site of the SH is assumed to mediate the efficient neutralization of O2 attacking the active site. This process is facilitated presumably by the extended redox capability provided through an additional FMN molecule bound to the hydrogenase module (Lauterbach et al. 2011b). Enzymes that are virtually insensitive to oxygen are represented by the subclass of regulatory H2-sensing [NiFe] hydrogenases such as those described for R. eutropha and Rhodobacter capsulatus. These sensory proteins are characterized by a turnover rate of H2 oxidation that is two orders of magnitude lower than the rate of MBH-like hydrogenases and the lack

of inactive states such as Niu-A and Nir-B. However, the proteins are rendered O2-sensitive when two bulky amino acids that likely limit the gas access to the active site are replaced by smaller residues (Buhrke et al. 2005; Duche´ et al. 2005). The [NiFe(Se)] hydrogenases are often referred to as ‘‘O2resistant’’ since they also do not exhibit the inactive Niu-A state. Nevertheless, H2 oxidation activity by this subclass of [NiFe] hydrogenase is rapidly and completely inhibited in the presence of trace amounts of O2. However, at low levels of O2, the [NiFe (Se)] hydrogenase from Desulfomicrobium baculatum retains partial H2-evolving activity of H2 production (Parkin et al. 2008).

Biogenesis of Hydrogenases The complex architecture of the active sites in [NiFe], [FeFe], and [Fe] hydrogenases raises the question of how these metal centers are assembled and incorporated into the apoenzymes. Specific auxiliary proteins, which control cofactor assembly, insertion, and coordinated folding of the respective protein, are usually involved in the biosynthesis of metalloenzymes. Inspection of the nucleotide sequences of hydrogenase gene clusters revealed numerous highly conserved accessory genes closely linked to the corresponding structural genes, and mutant analysis delivered the first evidence that the products of these genes participate in complex posttranslational metal insertion

H2-Metabolizing Prokaryotes

. Table 4.3 Characteristics of Hyp proteins

H2 ↔ 2e− + 2H+ O2 + 4e− + 4H+ → 2H2O

Sizea Protein (kDa) Domain(s)/motif(s)

e−

H2O

13.2

Ni-binding domain, Zn-binding domain (Watanabe et al. 2009)

Complex formation with HypB, Ni insertion

HypBc

31.6

GTPase domain, histidinerich region (Gasper et al. 2006)

Complex formation with HypA, Ni insertion, and in some cases, Ni storage

HypC

9.7

HypD

41.4

[4Fe-4S] cluster binding domain, putative nucleotide binding site (Watanabe et al. 2007)

Fe-dependent complex formation with HypC, scaffold complex

HypE

33.7

ATP-binding site (Watanabe et al. 2007)

ATP-dependent dehydration of thiocarboxamide yielding thiocyanate

HypF

82.0

ATP-dependent Zn-finger domain, carbamoylation of nucleotide-binding YrdCHypE like domain, acylphosphatase domain, Kae1-like universal domain (Petkun et al. 2011)

HypXc

65.7

N10-formyltetrahydrofolate binding site and enoyl-CoA hydratase/isomerase signature (Burgdorf et al. 2005)

O2

H2

O2

. Fig. 4.16 Proposed gas channel (blue arrows), water tunnel network (red arrow), and the electron relay (black arrow) in the MBH of R. eutropha. The catalytic model (see text) predicts H2O formation as a byproduct of H2 conversion in the presence of O2. The large (blue) and small (green) subunits are shown in ribbon representation with reduced opacity. Relevant cavities are shown in gray. Water molecules within these cavities are visualized as red spheres (Adapted from Fritsch et al. (2011b))

pathways. This chapter is devoted to the maturation of hydrogenases (reviewed by Forzi and Sawers 2007; Leach and Zamble 2007; McGlynn et al. 2009; Mulder et al. 2011; Nicolet and Fontecilla-Camps 2012) and does not address the acquisition of metals, the general machinery for protein folding, and the biogenesis of Fe-S clusters and other prosthetic groups commonly found in metalloproteins. Depending on the cellular location and physiological function, hydrogenases undergo various stages of maturation.

Maturation of [NiFe] Hydrogenases At least six hyp gene products (HypA, HypB, HypC, HypD, HypE, and HypF), present in bacterial as well as archaeal [NiFe] hydrogenase-containing species, are involved in the assembly and insertion of the heterodinuclear metal center (> Table 4.3). The designation hyp stands for ‘‘genes affecting hydrogenases pleiotropically’’ meaning that mutations in the individual hyp genes either reduce or abolish the activity of multiple [NiFe] hydrogenase isoenzymes, as demonstrated for Escherichia coli (Lutz et al. 1991), Ralstonia eutropha (Buhrke et al. 2001; Dernedde et al. 1996; Wolf et al. 1998), and Thiocapsa roseopersicina (Maro´ti et al. 2003).

Putative functionb

HypA

H2 Ni-Fe

4

OB fold domain, conserved Chaperone which N-terminal cysteine residue forms complexes (Watanabe et al. 2007) with either HypD or the large subunit

Maturation of the [NiFe] active site under aerobic conditions

a

Sizes of E. coli K12 Hyp proteins are given (Lutz et al. 1991; Maier and Bo¨ck 1996) b References are given in the text c HypB of E. coli is devoid of a polyhistidine sequence. HypX is not present in E. coli

The sequence of events leading to the formation of active [NiFe] hydrogenase is gradually emerging and takes advantage of crystal structures that are available now for all six Hyp proteins (Gasper et al. 2006; Watanabe et al. 2007, 2009; Petkun et al. 2011). The current model of the maturation pathway is predominantly based on studies on hydrogenase-3 (Hyd-3) of E. coli. Its [NiFe] center-containing subunit HycE is part of a protein complex attached to the inner side of the cytoplasmic membrane (Rossmann et al. 1994; > Fig. 4.2). The working model (> Fig. 4.17) postulates a sequential insertion of an Fe(CN )2CO moiety and the nickel ion into the HycE precursor. It has been suggested that the HypC-HypD complex acts as a scaffold for the assembly of the Fe(CN )2CO moiety (Bo¨ck et al. 2006; Watanabe et al. 2007). The CN ligand is synthesized

159

160

4

H2-Metabolizing Prokaryotes

Cys CysN1 B1

Fe CO ATP

O

AMP + PPi

HypC

?

CO

HypF

CO

Fe CN

CN

CN

CN

SH S

Fe

C OPO32− NH2

HypE

CysB2 CysN2

HypC

HypD ATP

SH S

ADP + Pi

Ni2+ Cys CysN1 B1

GTP

HypC

HypB

S

S CO

HypA

Ni

Fe CN CN

GDP + Pi

S

S

CysB2 CysN2

Specific Endopeptidase

Cys CysN1 B1 S Fe-S

S CO

Ni

Fe CN CN

S

S

CysB2 CysN2

. Fig. 4.17 Model for the maturation of standard [NiFe] hydrogenases based mainly on studies of E. coli hydrogenase-3. The large and small subunits of the hydrogenase are shown in dark blue and light blue, respectively. The Hyp proteins are depicted in green. Iron-sulfur clusters are indicated as diamonds (Fe-S). The sources of Fe and CO in the Fe(CN )2CO complex remain to be elucidated. See text for details

on a strictly conserved C-terminal cysteine of HypE via a HypFmediated process (Blokesch et al. 2002; Paschos et al. 2001; Reissmann et al. 2003). HypF, which contains several conserved functional domains (> Table 4.3; Petkun et al. 2011), converts carbamoylphosphate to carbamoyladenylate in an ATPdependent reaction. In a transient (HypE-HypF)2 complex (Rangarajan et al. 2008), HypF mediates the carbamoylation of the C-terminal cysteine of HypE. Finally, HypE catalyzes the ATP-dependent dehydration of the thiocarboxamide yielding thiocyanate and subsequently transfers the cyano group to the iron situated on HypC-HypD (Paschos et al. 2001; Blokesch et al. 2002, 2004; Watanabe et al. 2007). Scarcely anything is known about the origin of the CO group and how it gets positioned onto the iron atom. Labeling experiments exclude carbamoylphosphate as the source of CO (Forzi et al. 2007; Lenz et al. 2007). Externally supplied gaseous CO can serve as the carbonyl source, but ambient CO levels are too low to meet the

CO requirements as demonstrated for the regulatory hydrogenase from R. eutropha, leading to the conclusion that the carbonyl ligand originates rather from an intracellular metabolite than an external substance (Bu¨rstel et al. 2011). It is postulated that the Fe(CN )2CO group on the HypCHypD complex is directly transferred to the large HycE subunit of Hyd-3 from E. coli (Blokesch et al. 2002). This involves a transient complex formation of HycE with HypC via one of the nickel-coordinating cysteine residues (CysN1; > Fig. 4.17). In this process, disulfide bridge formation between HycE and HypC can be excluded (Magalon and Bo¨ck 2000). Due to the lack of data from other organisms, it is premature to generalize the maturation pathway. Indeed, genetic analysis suggests the existence of variations of a common principle. In Thiocapsa roseopersicina, two distinct HypC species are required for the assembly of the hydrogenases (Maro´ti et al. 2003). Several protein complexes including HypC-HypD, HypE-HypF1, HypC-HypD-HypE, and

H2-Metabolizing Prokaryotes

HypC-HoxH have been detected in R. eutropha (Jones et al. 2004). Moreover, maturation of periplasmically oriented membrane-bound [NiFe] hydrogenases depends on sets of additional accessory proteins (see below). In the second stage of cofactor assembly, nickel is incorporated by the concerted action of the HypA and HypB proteins (Olson et al. 2001; Hube et al. 2002; Chan et al. 2012; > Fig. 4.17). HypB proteins share a GTP-binding site, and GTP hydrolysis is essential for dimerization of HypB (Maier et al. 1995; Gasper et al. 2006). However, to some extent, the lack of hypA and hypB can be complemented by supplying high concentrations of Ni2+ to the medium (Waugh and Boxer 1986; Hube et al. 2002). Unlike E. coli HypB, the majority of HypB proteins contain polyhistidine stretches with Ni-chelating potential. In fact, in Bradyrhizobium japonicum HypB, this stretch is capable of binding 16 nickel ions per dimer (Fu et al. 1995). This observation suggests that in some organisms, HypB has a dual function, which is Ni storage and Ni delivery to hydrogenase (Dias et al. 2008). The peptidyl-prolyl cis/trans-isomerase SlyD also seems to be involved in the nickel insertion step. SlyD, that itself has a histidine-rich C-terminus, forms a complex with HypB and stimulates the release of Ni from the highaffinity Ni-binding site in HypB (Zhang et al. 2005; Leach and Zamble 2007). An additional Hyp protein, HypX (formerly designated ‘‘HoxX’’), has been found only in aerobic H2-oxidizing bacteria, including B. japonicum (Van Soom et al. 1993), R. eutropha (Lenz et al. 1994), and R. leguminosarum (Rey et al. 1996). Mutations in hypX decreased the activity of hydrogenase under certain growth conditions in two nitrogen-fixing organisms (Durmowicz and Maier 1997; Rey et al. 1996) as well as in a chemolithotroph (Wolf et al. 1998). The HypX protein reveals two interesting sequence motifs: an N-terminal N10-formyltetrahydrofolate binding site and a C-terminal signature that is typical for enoyl-CoA hydratases and isomerases. It was postulated that HypX plays a role in the recruitment of diatomic ligands in a tetrahydrofolate (THF)-coupled reaction (Rey et al. 1996) but its exact role is still a matter of discussion (Horch et al. 2012). Genomic sequencing has uncovered hypX homologs in other prokaryotes including Aquifex aeolicus (Deckert et al. 1998) and Streptomyces avermitilis (Omura et al. 2001). Once the [NiFe] center is incorporated into the large hydrogenase subunit (> Fig. 4.17), the chaperone HypC dissociates from the precursor. In the case of Hyd-3 from E. coli, the specific endopeptidase HycI completes the reaction by cleaving off 32 amino acids from the C-terminus of the HycE precursor (Rossmann et al. 1994). Supposedly, the endopeptidase to ‘‘inspects’’ correct nickel insertion and to trigger a conformational change by proteolysis, henceforth allowing the hydrogenase to enter a folded, oligomeric state. Crystal structure analysis of HybD, the endopeptidase specific for Hyd-2 from E. coli, has uncovered a metal binding site (Glu16, Asp62, His93) which is implicated in nickel recognition during the proofreading process (Fritsche et al. 1999; Theodoratou et al. 2000). In organisms containing multiple [NiFe] hydrogenases, C-terminal proteolysis is mediated by individual endopeptidases

4

which are not functionally exchangeable. The length of the cleaved peptides varies considerably (between 13 and 32 amino acids), whereas the cleavage site seems to be rather conserved. It consists of a basic amino acid separated by three residues from the terminal Ni-coordinating cysteine (CysB2; > Fig. 4.17). The exact cleavage position was experimentally determined to be an arginine residue in HycE of the E. coli Hyd-3 (Rossmann et al. 1994; Theodoratou et al. 2000). In the processed large subunit of the NAD+-reducing hydrogenase from R. eutropha, a histidine residue was found at the C-terminal position (Thiemermann et al. 1996) as also deduced from the crystal structures of periplasmic [NiFe] hydrogenases. Not all hydrogenases, however, undergo C-terminal proteolysis during maturation. The H2-sensing proteins (reviewed by Vignais and Billoud 2007), the CO-induced hydrogenase of Rhodospirillum rubrum (Fox et al. 1996a, b), and the Ech hydrogenase of Methanosarcina barkeri (Ku¨nkel et al. 1998) are welldocumented examples. Although experimental results have shown that metal center insertion into the H2-sensing proteins of B. japonicum (Olson et al. 1997), R. capsulatus (Colbeau et al. 1998), and R. eutropha (Buhrke et al. 2001) relies on hyp gene products, a C-terminal extension at the active site subunit is absent and dispensable for metal center assembly. On the other hand, it was unambiguously demonstrated for hydrogenases such as Hyd-3 of E. coli (Binder et al. 1996) and the NAD+reducing hydrogenase of R. eutropha (Massanz et al. 1997) that the C-terminal extension of the catalytic subunit is absolutely necessary for biogenesis of active enzyme.

Membrane Translocation of Hydrogenases Periplasmic and periplasmically oriented membrane-bound [NiFe] and [FeFe] hydrogenases are exported in the folded, cofactor-containing state across the cytoplasmic membrane. This transfer is mediated through the Tat translocon (reviewed by Fro¨bel et al. 2012). Proteins that undergo this mode of translocation share an N-terminal signal peptide, referred to as the ‘‘twin-arginine leader,’’ bearing a conserved consensus (S/T) RRxFLK motif (where x is a polar amino acid). In Tat deficient mutants of Azotobacter chroococcum (Yates et al. 1997), E. coli (Rodrigue et al. 1999), and R. eutropha (Bernhard et al. 2000), membrane targeting of [NiFe] hydrogenases is disrupted, and the proteins accumulate in the cytoplasm. Although the cytoplasmic enzymes are physiologically inactive, they usually exhibit high H2 oxidation activity which clearly shows that metal center assembly by the Hyp proteins occurs in the cytoplasm prior to translocation of the hydrogenase. Only the small subunit of [NiFe] hydrogenases carries the N-terminal twin-arginine leader peptide of 30–50 amino acid residues, whereas the large subunit lacks an export signal. Early observations on the [FeFe] hydrogenase of Desulfovibrio vulgaris Hildenborough led to the proposal that the large subunit is cotranslocated with the small subunit (van Dongen et al. 1988). Experimental evidence has accumulated which clearly supports a tandem export of the two

161

162

4

H2-Metabolizing Prokaryotes

subunits. Studies conducted with [NiFe] hydrogenases from E. coli (Menon et al. 1991; Menon and Robson 1994; Rodrigue et al. 1999), Desulfovibrio vulgaris (Nivie`re et al. 1992), R. eutropha (Bernhard et al. 2000), Desulfovibrio gigas (Rousset et al. 1998a), and Wolinella succinogenes (Gross et al. 1999) are all in line with the cotranslocation model. Crystal structure analysis of the [FeFe] hydrogenase from Desulfovibrio desulfuricans (Nicolet et al. 1999) has confirmed earlier predictions concerning export of this protein (Voordouw and Brenner 1985). In the mature functional molecule, the 34-amino-acid twin-arginine signal peptide, deduced from the DNA sequence of the small subunit, is missing in the crystallized protein (Hatchikian et al. 1999; Nicolet et al. 1999). This observation clearly indicates that periplasmic [FeFe] hydrogenases undergo a similar translocation process as [NiFe] hydrogenases. A second C-terminal peptide of 24 amino acids at the large

hoxK G

HoxK

E CO CN HoxV CN

F1 B1 A1

Fe(CN−)

Ni

Cys366

2CO

O2

D

V

Isc/Suf

C

R

HypC

HypD

HoxL ATP

X

L

S

Fe HypE

E

Metallocenter assembly

HoxG

Fe

CO CN CN

C OPO32−NH2

MBH-specific maturation

S

HypC

?

HypF1

It seems to be a common feature that membrane-bound [NiFe] hydrogenase operons contain, in addition to the hyp genes, sets of tightly linked accessory genes. The periplasmically oriented membrane-bound [NiFe] hydrogenase (MBH) of R. eutropha undergoes a particularly complex maturation process as depicted in > Fig. 4.18. The megaplasmid pHG1-encoded gene

Cys52

Fe CO

O

Additional Functions Involved in the Maturation of Periplasmically Oriented Membrane-Bound [NiFe] Hydrogenases

Z M LO QRT V hypA1B1 F1 C D

MBH structural genes

ATPAMP + PPi

subunit of D. desulfuricans [FeFe] hydrogenase is also absent in the mature protein. It remains to be elucidated whether this modification is related to translocation or other maturation functions.

Q

Fe Ni

ADP + Pi Cys Cys75 78

HoxL

S H

S

H S S

Fe

O

M CO CN CN

Transient super complex HoxG Fe

Cys597 Cys600

Ni

O2

Ni2+

T

Cytoplasm

Tat

Cys78

Cys75

GTP HypB1 GDP + Pi

HoxL HypA1

S S

Ni Fe S

CO CN CN

2e− HoxG

Periplasm

Cytb Fe Fe

S

Cys597 Cys600

Fe Ni

H2 2H+

. Fig. 4.18 Arrangement of genes coding for the membrane-bound [NiFe] hydrogenase (MBH) of R. eutropha and factors involved in the maturation of this O2-tolerant enzyme (Adapted from Fritsch et al. (2011a)). The genes hoxKGZ encode the small and large hydrogenase subunits (blue) and a membrane-integral b-type cytochrome (gray); the hoxMLOQRTV genes (yellow) code for MBH-specific accessory proteins; the hypA1B1F1CDEX genes (green) are responsible for [NiFe] site assembly (for the sake of simplicity, downstream genes involved in the regulation of hydrogenase gene expression are not shown). HoxR and HoxT support maturation at high O2 concentrations. Isc and Suf symbolize the general machinery for assembly and insertion of Fe-S clusters and are likely involved in incorporation of the Fe-S centers into HoxK. Folding and oligomerization of the HoxG subunit of MBH are triggered by the specific endoprotease HoxM (Bernhard et al. 1996), which was also shown to have a proofreading function (Magalon et al. 2001). See the text for further details and references

H2-Metabolizing Prokaryotes

cluster required for active MBH expression in R. eutropha encompasses 21 genes (> Fig. 4.18; Schwartz et al. 2003). Only three of these code for the structural polypeptides of the enzyme. Assembly and incorporation of the [NiFe] cofactor is accomplished by the products of at least six hyp genes (Jones et al. 2004). Moreover, a specific chaperone, HoxL, and a transfer protein, HoxV, which is likely involved in shuttling or storing the Fe(CN )2CO cofactor intermediate (> Fig. 4.18), are required for MBH maturation in R. eutropha (Ludwig et al. 2009). The need for this additional transfer step in maturation of MBH is not self-evident; however, HoxL and HoxV may play a protective role in the O2-exposed maturation of the large subunit. Sensitivity of the large subunit precursor toward oxygen was deduced from in vitro studies on Hyd-2 from E. coli conducted with cellfree extracts (Soboh et al. 2010). Mutant studies provided the first insights into the function of the accessory proteins which are not completely conserved in the various species. Inactivation of the accessory genes hyaE and hyaF in the hydrogenase-1 operon of E. coli (Menon et al. 1991) and the homologous genes hoxO and hoxQ in the MBH operon of R. eutropha (Bernhard et al. 1996) yields catalytically inactive hydrogenase, which accumulates in the cytosol. A dual function has been assigned to the proteins HoxO and HoxQ. They interact with the small subunit of R. eutropha via the twin-arginine leader peptide, thereby preventing premature export of the MBH. Furthermore, it has been suggested that the two proteins are suggested to protect the Fe-S clusters against reactive oxygen species (> Fig. 4.18; Schubert et al. 2007). Homologous clusters of hoxO, hoxQ, hoxR, and hoxT genes are found predominantly in knallgas bacteria. Investigations with mutant stains of R. eutropha demonstrated that HoxR and HoxT are key components in MBH maturation at ambient O2 levels whereas both proteins are dispensable when cells are grown under O2-limited conditions (> Fig. 4.18; Fritsch et al. 2011a). Moreover, it was suggested that the rubredoxin-like HoxR protein has a specific function in the biogenesis of the unusual [4Fe-3S] center in MBH under aerobic conditions (Fritsch et al. 2011a). A hydrogenase-specific chaperone function was assigned to the HoxT homolog HybE in E. coli where it presumably prevents premature targeting and translocation of Hyd-2 (Jack et al. 2004).

Maturation of [FeFe] Hydrogenases Compared to [NiFe] hydrogenases, biosynthesis of the H-cluster of [FeFe] hydrogenase appears to be less complex, involving only three specific accessory genes (reviewed by Mulder et al. 2011; Nicolet and Fontecilla-Camps 2012). These genes, designated hydE, hydF, and hydG, were initially discovered in screens of C. reinhardtii mutants incapable of H2 production (Posewitz et al. 2004a, b). The HydE and HydG proteins belong to the radical S-adenosylmethionine (radical SAM) superfamily of enzymes containing the canonical CX3CX2C sequence motifs. Each of the two proteins from Thermotoga maritima carries a [4Fe-4S] cluster bound to the CX3CX2C sequence (Rubach

4

et al. 2005). In addition, reconstituted HydE from T. maritima contains a [2Fe-2S] cluster that is not essential for production of active [FeFe] hydrogenase (Nicolet et al. 2008). Some green algae encode a gene fusion of hydE and hydF (Bo¨ck et al. 2006). HydF has an N-terminal GTPase domain comprised of the Walker A P-loop, Walker B Mg2+-binding motifs, and a putative dimerization domain (Cendron et al. 2011). The C-terminal domain of HydF contains highly conserved cysteine and histidine residues. Reconstituted HydF from T. maritima was found to hydrolyze GTP to GDP and to contain a single [4Fe-4S] cluster (Brazzolotto et al. 2006). Amino acid substitution studies demonstrated that the highly conserved radical SAM domains of both HydE and HydG as well as the GTPase domain and Fe-S binding motifs of HydF are essential for the maturation of the H-cluster-containing HydA protein (King et al. 2006). HydA protein heterologously expressed in a genetic background devoid of HydE, HydF, and HydG (HydADEFG) could be activated by addition of a cell extract containing coexpressed HydE, HydF, and HydG (McGlynn et al. 2007). HydF purified from extracts containing HydG and HydE is able to activate HydADEFG (McGlynn et al. 2008). Biochemical and spectroscopic analysis of this form of HypF revealed the presence of a [4Fe-4S] cluster in addition to a [2Fe-2S] moiety and CO as well as CN ligands (Shepard et al. 2010b). This experimental evidence provided the basis for the development of a hypothetical pathway of H-cluster biosynthesis (> Fig. 4.19). According to this model, HydE and HydG are responsible for the synthesis of the H-cluster precursor on a [2Fe-2S] cluster framework in the scaffold HydF. Subsequently, HydF transmits the 2Fe subcluster to the apo form of HydA. The crystal structure of HydADEFG protein from Clamydomonas reinhardtii heterologously expressed in E. coli provided important clues on the process of precursor transfer from HydF to HydA (Mulder et al. 2010). The structural analysis revealed that the [4Fe-4S] cubane of the H-cluster is already present at the active site of HydADEFG. This indicates that, at least in this experimental design, the cubane is assembled by the general Fe-S cluster biosynthetic pathway before the 2Fe subcluster is transferred from HydF to HydA (> Fig. 4.19). It is anticipated that the subcluster is inserted through a positively charged channel that closes subsequently via conformational changes in two conserved loop regions (Mulder et al. 2010). The exact role of GTP binding and hydrolysis of HydF is still unclear, but the functions appear to be relevant for precursor formation via action of HydE and HydG rather than for the 2Fe subcluster transfer process (Mulder et al. 2011). It has been shown that both CN and CO ligands are produced by HydG-catalyzed tyrosine degradation that presumably involves oxidative decarboxylation of dihydroglycine (Driesener et al. 2010; Shepard et al. 2010a). Interestingly, CO synthesis may depend on a second [4Fe-4S] cluster in HydG (Shepard et al. 2010a; Tron et al. 2011). The precise role of HydE in the biosynthetic pathway is still an unresolved issue. Based on available biochemical and spectroscopic data of the other two maturases HydG and HydF, HydE is considered to be responsible for production of the bridging dithiolate ligand.

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H2-Metabolizing Prokaryotes

CO and CNbiosynthesis

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Dimethylamine biosynthesis

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2H+ + 2e−

H2

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Fe

Fe

Fe

S

NC

C O

CN

Fe CO

HydF[4Fe-4S]

OC

NC

Fe C O

CN

Cys CO

HydA

. Fig. 4.19 Model for the [FeFe] hydrogenase (HydA) maturation pathway. See text for details

The carbon-based substrate of this reaction is yet unknown. The discussion is focused on a common metabolite since coexpression of the three maturases and HydA in E. coli, that lacks [FeFe] hydrogenase, leads to production of active enzyme (McGlynn et al. 2007). The identified hydEF gene fusions and copurification of HydE with HydF under certain conditions suggest a concerted action of the two proteins (McGlynn et al. 2008).

Maturation of [Fe] Hydrogenases At present, there is very little experimental data available on the events leading to the insertion of the Fe-GP cofactor into [Fe] hydrogenase. Genome analysis of several methanogenic archaea identified seven genes hcgABCDEFG (hmd co-occurring genes) that are usually clustered with the [Fe] hydrogenase structural gene hmd (Thauer et al. 2010). hcgA has the coding capacity for a protein similar to biotin synthases and other radical SAM enzymes. However, it lacks the N-terminal CX3CX2C or CX4CX2C motif characteristic for the radical SAM protein family. This motif coordinates the [4Fe-4S] cluster that is essential for radical formation (Duffus et al. 2012). Instead, the hypothetical hcgA gene product harbors a conserved CX5CX2C motif (McGlynn et al. 2010). According to the crystal structure of HcgB of Methanobacterium thermoautotrophicus (PDB entry: 3BRC), the homodimeric protein shares structural similarities with a human inosine triphosphatase, which catalyzes the cleavage of pyrophosphate from ITP (Schick et al. 2012). Moreover, it was postulated that the hcgC gene product contains a putative NAD (P)-binding domain, and the putative hcgD-encoded protein reveals a sequence similar to a protein in yeast that interacts with the transcriptional activator NGG1p. Finally, the hcgE gene product shares similarity to proteins catalyzing ATP-dependent ubiquitin activation whereas the gene hcgF could not be related

to any product of known function (Thauer et al. 2010). More recently, the gene hcgG that was originally annotated as a gene for a fibrillarin-like protein was suggested to encode a SAMdependent methyltransferase (Schick et al. 2012). The Fe-GP cofactor can be reversibly extracted from [Fe] hydrogenase under mildly acidic or alkaline conditions in the presence of mercaptoethanol (Shima et al. 2012). In order to obtain first insights into Fe-GP cofactor biosynthesis, in vivo labeling experiments with 13C- and 2H-labeled precursors were carried out with Methanothermobacter marburgensis and Methanobrevibacter smithii, and the extracted cofactor was analyzed by NMR, mass spectrometry, and infrared spectroscopy (Schick et al. 2012). The labeling pattern was found to be consistent with a synthesis of the Fe-GP cofactor from one acetate, two pyruvate, three CO2, and a methyl group of methionine (> Fig. 4.20). The biosynthesis of the pyridinol moiety was found to be distinct from that of pyridine polyketides. Remarkably, also the biosynthesis of the CO ligands at the iron of the Fe-GP cofactor appears to be different from that of [FeFe] hydrogenases, since dehydroglycine could be excluded as a biosynthetic precursor. Schick et al. (2012) postulated that one methyl group attached to the pyridinol ring could be introduced by HcgG in a SAM-dependent reaction and that the HcgB may catalyze the ligation of the GMP and pyridinol parts of the Fe-GP cofactor. The labeling pattern of GMP follows the canonical biosynthetic pathway.

Genetic Organization [NiFe] Hydrogenases The bulk of the presently available nucleotide sequence data relates to the [NiFe] hydrogenases. Sorting these sequences solely on the basis of operon structure reveals groups with more or less conserved gene patterns. On the one hand, these

4

H2-Metabolizing Prokaryotes

OC CO2

°

Fe2+ C O

3 acetate

O

S

M. mazei vho

* °CH -COO -

Cys

N *

HO

(-S-CH3)

G

* °

* # #

4500 nt

O

GMP

A

C

A

C

M. mazei vht 4500 nt

#

CH3-CO-COO-

G

D

pyruvate

L-methionine 500 bp

. Fig. 4.20 Putative biosynthetic precursors of the iron-guanylylpyridinol cofactor of [Fe] hydrogenase of Methanothermobacter marburgensis and Methanobrevibacter smithii as deduced from 13C labeling experiments (Adapted from Schick et al. (2012)). See text for details

groups correlate to a certain extent with the subclusters defined by clustering analysis of the deduced amino acid sequences of the hydrogenase subunits (Vignais et al. 2001; Vignais and Billoud 2007). On the other hand, they reflect common biochemical and physiological properties of the respective enzymes.

Archaeal Membrane-Bound Hydrogenases In Methanosarcina mazei, two operons (vhoGAC and vhtGACD) encoding F420-nonreactive hydrogenases have been identified and sequenced (Deppenmeier 1995; Deppenmeier et al. 1995). Each operon contains genes for the hydrogenase small (vhoG and vhtG) and large subunits (vhoA and vhtA) followed by a gene for a b-type cytochrome (vhoC and vhtC). The vht operon contains a fourth gene, vhtD, which predicts a protein with similarity to the maturation proteases associated with bacterial membrane-bound hydrogenases, suggesting that at least one of the two M. mazei F420-nonreactive hydrogenases undergo a similar maturation process. The genomic sequence of M. mazei provides additional evidence for this assumption (Deppenmeier et al. 2002): A set of hyp genes (hypC, hypD, hypA, and hypE) is located adjacent to the vho operon in the opposite orientation. Furthermore, tandem copies of hypB and hypC are found a few kilobases away from the vhtGACD operon, and solitary copies of hypF and hypE are present at remote sites. Northern analysis confirmed that the vho and vht operons are both expressed as single transcripts approximately 4,500 nucleotides (nt) in size (> Fig. 4.21).

Bacterial Membrane-Bound Hydrogenases The closest relatives of the Vho and Vht hydrogenases of M. mazei are the bacterial membrane-bound hydrogenases. Examples of this type of hydrogenase are found in both Gram-negative and Gram-positive bacteria. The most thoroughly studied representatives are the membrane-bound hydrogenases of various proteobacteria (Friedrich and Schwartz

. Fig. 4.21 Organization of the vho and vht operons of Methanosarcina mazei. Light brown arrows represent genes for the hydrogenase small subunits, dark brown arrows, genes for the large subunits, and orange arrows, genes for b-type cytochromes. A gene for a putative maturation protease, vhtD, is shown in gray. The blue arrows symbolize experimentally determined transcripts (sizes given in nucleotides)

1993; Vignais and Toussaint 1994). In contrast to the vho and vht operons, the proteobacterial genes form large, polycistronic transcriptional units. In the following, four examples of this group are discussed: the MBH locus of Ralstonia eutropha, the hup/hyp regions of Rhodobacter capsulatus and Rhizobium leguminosarum, and the Escherichia coli hya operon. The MBH locus of R. eutropha is a contiguous series of hydrogenase genes occupying approximately 23 kb on the megaplasmid pHG1 (Kortlu¨ke et al. 1992; Dernedde et al. 1993, 1996 > Fig. 4.22). The first two genes, hoxK and hoxG, which code for the small and large subunit of the hydrogenase, respectively, are followed by a gene (hoxZ) encoding a b-type cytochrome. This is the primary electron acceptor of the membrane-bound hydrogenase and also mediates attachment of the enzyme to the outer surface of the cytoplasmic membrane (Bernhard et al. 1997). The hoxZ gene is followed by a series of MBH-specific accessory genes designated M, L, O, Q, R, T, and V. The hoxM gene encodes a specific protease required for the C-terminal processing of the large subunit. The function of the other gene products is discussed above in the section > ‘‘Maturation of [NiFe] Hydrogenases.’’ Immediately downstream of hoxV lies a second group of accessory genes called ‘‘hyp genes.’’ The hyp genes are involved in the assembly of the [NiFe] center of the hydrogenase active site (see the section > ‘‘Maturation of [NiFe] Hydrogenases’’). Immediately downstream of the R. eutropha hyp region is a set of regulatory genes designated ‘‘hoxA,’’ ‘‘hoxB,’’ ‘‘hoxC,’’ and ‘‘hoxJ’’ (Lenz et al. 1997). Their function will be discussed below in detail. The same basic pattern of genes with minor variations is found in several other representatives of the proteobacteria. In R. capsulatus, a hypF gene is absent from the hyp operon. In its place is the regulatory gene hupR1. A set of regulatory genes (hupTUV), which corresponds to hoxJ/hoxB/hoxC, is encoded together with a hypF gene in a transcriptional unit immediately upstream of the main hydrogenase operon (Elsen et al. 1996; > Fig. 4.22). In R. leguminosarum, an additional gene (hupE), encoding a Ni transporter of the HupE/UreJ protein family

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H2-Metabolizing Prokaryotes

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E. coli A

hyp B A

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C D

F E

. Fig. 4.22 Organization of hydrogenase operons of R. eutropha, R. capsulatus, R. leguminosarum, and E. coli. Color-coding emphasizes similarity at the level of deduced amino acid sequences. Light brown arrows represent genes for the small subunits, dark brown arrows, genes for the large subunits, and orange arrows, genes for b-type cytochromes. Genes for maturation proteases are shown in gray. The R. hyp genes are shown in shades of yellow and gold and other accessory genes, in shades of blue. NtrC-type response regulators are black, and histidine protein kinases are red

(Eitinger et al. 2005; Brito et al. 2010), is located downstream of the protease gene, hupD. In Methylococcus capsulatus (Bath), hupE is adjacent to the structural genes hupSL (Csa´ki et al. 2001). The E. coli hya operon, which encodes a membranebound hydrogenase isoenzyme designated ‘‘hydrogenase-1,’’ is simpler than the three examples discussed above (Menon et al. 1990; > Fig. 4.22). This is partly due to the fact that maturation of this enzyme is mediated by the Hyp proteins encoded in an operon adjacent to the hyc operon. In addition to a gene for a maturation protease, hyaD, two other accessory genes, hyaE and hyaF, are present. It is not clear whether, aside from the Hyp proteins, accessory proteins encoded in the other hydrogenase operons are required for the function of hydrogenase-1. An interesting variant is the hynSL operon of Thiocapsa roseopersicina. The genes hynS and hynL, which encode the small and large subunits, respectively, of a membrane-bound hydrogenase, are separated by two reading frames designated isp1 and isp2. These reading frames are not related to any known hydrogenase accessory genes (Ra´khely et al. 1998).

The Cytoplasmic F420-Nonreactive Hydrogenases The operons encoding the F420-nonreactive hydrogenases of Methanothermobacter thermoautotrophicus (mvhDGAB) and

M. voltae (vhcDGAB and vhuDGAB; Reeve et al. 1989; Halboth and Klein 1992) are representatives of another conserved family (> Fig. 4.23). The first open reading frame (ORF) of each operon predicts a protein of unknown function, followed by genes for the small (mvhG, vhcG, and vhuG) and large subunits (mvhA, vhcA, and vhuA) of the hydrogenase. A fourth gene encodes a polyferredoxin, which may be the primary electron acceptor interacting with the hydrogenase in vivo. The vhuDGAUB operon is an interesting variant (Halboth and Klein 1992). In place of a gene for the large subunit, as is found in the other operons, there are two ORFs. The second, short ORF, designated vhuU, predicts a polypeptide related to the C-terminus of typical large subunits. This is particularly intriguing since this part of the protein participates in coordination of the [NiFe] center.

The F420-Reactive Hydrogenases Genes for F420-reactive hydrogenases have also been cloned and sequenced. Methanococcus voltae and M. barkeri each contains two such operons (Halboth and Klein 1992; Vaupel and Thauer 1998; > Fig. 4.24). The four operons share the same pattern: The hydrogenase large and small subunits are encoded by the first and third genes, respectively. The final gene of the set encodes

4

H2-Metabolizing Prokaryotes

M. thermoautotrophicus D

G

mvh B

A

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hox

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60

M. barkeri

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46

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fru

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56 46 5 5 46

. Fig. 4.23 Organization of the M. thermoautotrophicus mvh operon and of the M. voltae vhc and vhu operons. The genes for polyferredoxins are shown in pink. For additional details, see the legends to > Figs. 4.21 and > 4.22

. Fig. 4.25 Organization of the R. eutropha and R. opacus bidirectional hydrogenase loci and of the Synecchococcus PCC6301 (A. nidulans) and A. variabilis hox regions. The genes encoding subunits of the NADH oxidoreductase module are shown in shades of green and the hyp genes, in shades of yellow and gold. For additional details, see the legends to > Figs. 4.21 and > 4.22

frh 3600 nt

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D

G

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fre 3600 nt

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G

B

500 bp

E

. Fig. 4.24 Organization of the M. voltae fru operon and of the M. barkeri frh and fre operons. The genes for the subunit with the F420-binding site are dark red. For additional details, see the legends to > Figs. 4.21 and > 4.22

the third subunit of the trimeric enzyme, which may carry the site of interaction with the cofactor F420. In all four operons, the genes for the large and small subunits are separated by an additional ORF which probably encodes a specific protease responsible for the C-terminal proteolytic processing of the large subunit.

The Cytoplasmic, NAD-Reducing Hydrogenases The cytoplasmic, NAD-reducing hydrogenase (SH) of R. eutropha is the prototype of a family of multimeric enzymes related both structurally and physiologically. The hydrogenases of this family are often called ‘‘bidirectional.’’ The R. eutropha enzyme is encoded by a complex operon consisting of five genes for the catalytic subunits and four accessory genes (Tran-Betcke et al. 1990; Thiemermann et al. 1996; Wolf et al. 1998; > Fig. 4.25). The operon is expressed as a 7,600-nt primary

transcript which is apparently cleaved into smaller secondary mRNAs (Oelmu¨ller et al. 1990). The first two genes of the operon (hoxF and hoxU) code for the NADH oxidoreductase (diaphorase) moiety of the enzyme. The two genes immediately downstream (hoxY and hoxH) encode the small and large subunits, respectively, of the hydrogenase module. The product of hoxW is a specific protease which mediates C-terminal processing of the large subunit (Thiemermann et al. 1996). The product of hoxI is identical to the so-called B protein that contains a putative nucleotide binding site, similar to that of cAMP receptors. HoxI is coexpressed with the SH (Ka¨rst et al. 1987) and forms a complex with the diaphorase moiety (Burgdorf et al. 2005). Downstream of hoxI is a duplicated set of hyp genes: hypA2, -B2, and -F2 (Wolf et al. 1998). The operon encoding the soluble, hydrogenase of the Gram-positive bacterium Rhodococcus opacus seems to be a carbon copy of the R. eutropha operon. The sequenced segment of the R. opacus operon revealed the genes hoxF, -U, -Y, H, -W, and part of hoxI (Grzeszik et al. 1997a). A 7,500-nt transcript has been reported, suggesting that here, too, hyp genes are included in the hydrogenase mRNA. A similar set of genes directs the expression of a so-called bidirectional hydrogenase in unicellular cyanobacteria such as Synecchococcus PCC6301 (Anacystis nidulans) (Boison et al. 1996; Tamagnini et al. 2007). As in the case of R. eutropha, a set of structural genes (hoxUYH) is followed by a contiguous set of accessory genes (hoxWhypABF). A hoxI-like gene is missing in this series of genes. A hoxF homolog accompanied by a gene designated ‘‘hoxE’’ is encoded in a separate transcriptional unit located at a distance of 16 kb. HoxE is related to the protein NuoE of complex I and is a component of the mature hydrogenase

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(Schmitz et al. 2002). Another variant of the same genetic pattern is found in the filamentous cyanobacterium Anabaena variabilis. The bidirectional hydrogenase of this organism is encoded by an operon containing the genes hoxF, U, -Y, and -H (Schmitz et al. 1995). ORF8 and ORF3, which flank hoxY, code for proteins of unknown function (> Fig. 4.25).

Multimeric H2-Evolving Hydrogenases Both archaea and bacteria contain multimeric H2-evolving hydrogenases encoded in complex operons (> Fig. 4.26). The prototype of this class of enzymes is the E. coli hydrogenase-3, which uses electrons from formate to reduce protons, thereby generating H2. Hydrogenase-3 is encoded in a complex locus together with a set of Hyp proteins. The corresponding operons (hycBCDEFGHI and hypABCDE) are adjacent to each other and are transcribed from divergent promoters (Bo¨hm et al. 1990; > Figs. 4.26 and > 4.30). The gene for the positive regulator FhlA is located downstream of the hyp operon and is transcribed from its own promoter. Two additional genes form a separate transcriptional unit located in the same gene cluster: hypF, which encodes a maturation protein, and hydN. The product of hydN may have a function in electron transfer from or to formate dehydrogenase (Maier et al. 1996b). The hydrogenase large and small subunits are encoded by the genes hycE and hycG, respectively. The hycI gene encodes a specific endopeptidase. The function of the other gene products is largely unknown. The product of hycF is probably an Fe-S-containing protein. The hycC and hycD genes encode membrane-spanning proteins.

E. coli A

hyc

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D

E

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H I

The latter three proteins, as well as HycE and HycG, are related to components of the NADH-ubiquinone oxidoreductase complex (complex I) of E. coli and other organisms. Similar sets of genes have been identified in the anoxygenic phototroph Rhodospirillum rubrum (Fox et al. 1996a), the chemolithoautotroph Carboxydothermus hydrogenoformans (Soboh et al. 2002), and the methanogen M. barkeri (Ku¨nkel et al. 1998). The genomes of Pyrococcus furiosus, Methanothermobacter marburgensis, M. thermoautotrophicus, M. kandleri, and Methanocaldococcus janaschii encode related hydrogenases with a more complex subunit composition (Tersteegen and Hedderich 1999). E. coli contains a second operon of the same type designated ‘‘hyf ’’ (Andrews et al. 1997; Skibinski et al. 2002) that is not expressed at a significant level in the wild type (Self et al. 2004) and of unknown function. The operons of this family differ in gene number and order. Nevertheless, they all encode multisubunit hydrogenases containing proteins related to various components of complex I.

Putative High-Affinity Hydrogenases Representatives of this type of hydrogenase have been identified on the basis of sequence data in various taxonomic groups. Several strains of Streptomyces and Mycobacterium contain the corresponding genes. Related genes have also been reported for Gram-negative bacteria (Schwartz et al. 2003). The gene clusters typically contain genes for the small and large catalytic subunits separated from a set of hyp genes by four ORFs for proteins of unknown function (Constant et al. 2011). An example for this arrangement is found in Streptomyces avermitilis (> Fig. 4.27). The above summary leads to the conclusion that typical patterns of gene organization are correlated with the major structural types of hydrogenases. Thus, conserved metabolic and structural specialization are reflected in conserved operonic structures. Basic blueprints of genetic organization persist across taxonomic boundaries.

coo M

K

L X

Streptomyces avermitilis M. barkeri

ech

hyp

hhy

hup

hyp

6000 nt A

B

C D

E

X

F 1000 bp

. Fig. 4.26 Organization of the E. coli hyc operon, the R. rubrum coo operon, and the M. barkeri ech operon. Genes encoding proteins related to components of NADH-ubiquinone oxidoreductase I of E. coli are colored gray, orange, yellow and red, respectively. The gene products of E. coli hycC, hycD, hycE (partial), and hycF (and of the related genes) are isologous to E. coli proteins NuoL/NuoM/NuoN, NuoH, and NuoI, respectively. For additional details, see the legends to > Figs. 4.21 and > 4.22

S

L

1 2 3

4

D

B A

F

D

E

C 1000 bp

. Fig. 4.27 Organization of the Streptomyces avermitilis MH-4680 hhy-hyp region (nomenclature based on Constant et al. (2011)). The hhyS and hhyL genes encoding the small and large hydrogenase subunits, respectively, are shown in brown and the hyp genes in shades of yellow and gold. Conserved ORFs encoding proteins of unknown function are numbered. For additional details, see the legends to > Figs. 4.21 and > 4.22

H2-Metabolizing Prokaryotes

[FeFe] Hydrogenases The genetic information for [FeFe] hydrogenases is contained, judging by the available sequence data, in simple transcriptional units consisting of 1–3 genes. The [FeFe] hydrogenase of Desulfovibrio vulgaris (Hildenborough), the first hydrogenase ever to be cloned and sequenced, is encoded by a pair of genes for the large and small subunits (Voordouw and Brenner 1985). The structure of the hydAB operon of the closely related strain Desulfomicrobium norvegicum (formerly Desulfovibrio desulfuricans strain Norway) is the same (Hatchikian et al. 1999). The monomeric hydrogenase (CpI) of Clostridium pasteurianum is encoded by a solitary gene which appears to be a fusion of the paired genes which normally code for the two hydrogenase subunits (Meyer and Gagnon 1991; > Fig. 4.10). The same applies to the monomeric hydrogenase of M. elsdenii (Atta and Meyer 2000). The set of genes encoding the [FeFe] hydrogenase of the hyperthermophile Thermotoga maritima is a singular case. Three genes, designated ‘‘hydA,’’ ‘‘hydB,’’ and ‘‘hydC,’’ determine the a, b, and g subunits of the trimeric enzyme (Verhagen et al. 1999). The hydA gene encodes the basic hydrogenase moiety and resembles the ‘‘fused’’ gene of C. pasteurianum (> Fig. 4.10). The sequence of hydB predicts an Fe-S-containing flavoprotein related to HndC of D. fructosovorans. The deduced sequence of HydC reveals that this protein is related to NuoE of complex I. Owing to the lack of transcript data on the one hand and to the paucity of sequence data on the other, it cannot be ruled out that additional genes belong to the same transcriptional unit. To date, three [FeFe] hydrogenase accessory genes, designated hydE, hydF, and hydG, could be identified. These were initially discovered in screens of Chlamydomonas reinhardtii mutants incapable of H2 production (Posewitz et al. 2005). Homologs to all three genes can be found in all active [FeFe] hydrogenase-expressing organisms, whereas some green algae encode a gene fusion of hydE and hydF (Bo¨ck et al. 2006).

[Fe] Hydrogenases The [Fe] hydrogenases of M. marburgensis and other methanogenic archaea are N5,N10-methylene-H2MPT dehydrogenases (Afting et al. 1998). Genome analysis of several methanogenic archaea identified seven genes hcgABCDEFG (hmd co-occurring genes) that are clustered with the hmd gene for the hydrogenase catalytic subunit (reviewed by Thauer et al. 2010). For details on the functions of the corresponding gene products see the section on > ‘‘Maturation of [Fe] Hydrogenases’’.

Regulation of Hydrogenase Genes [NiFe] Hydrogenases The R. eutropha hox regulon is composed of two operons located on the 445-kb pHG1 megaplasmid and separated by ca. 70 kb

4

(Eberz et al. 1986; > Figs. 4.22 and > 4.25). The MBH operon encodes the heterotrimeric, membrane-bound hydrogenase and is transcribed as a >17-kb mRNA (Schwartz et al. 1999). The genes for the cytoplasmic, NAD-reducing hydrogenase belong to the SH operon. A 7,600-nt transcript has been mapped to this locus (Oelmu¨ller et al. 1990). Expression of the two operons is coordinate and responds to two environmental conditions (> Fig. 4.28a): (1) the availability of H2 and (2) the quality of the carbon and energy sources present in addition to H2 (Friedrich et al. 1981a; Friedrich 1982). The expression of the hydrogenase regulon is controlled at the level of transcription. The 24/ 12 promoters have been mapped upstream of the two operons, and their activity is strictly dependent on the minor transcription factor sN (Schwartz et al. 1998). The activity of PMBH and PSH is governed by the NtrC-type activator protein HoxA (Eberz and Friedrich 1991; Schwartz et al. 1998). This positive transcriptional regulator is encoded by the gene hoxA located downstream of the hyp genes in the MBH operon (Eberz and Friedrich 1991). A moderate, sD-dependent promoter (PhoxA) drives low-level, constitutive transcription of hoxA and perhaps of the downstream genes hoxB, hoxC, and hoxJ as well. Constitutive transcription of hoxA guarantees that at least a basal level of HoxA is present in the cell at all times, making sure that the organism is poised to respond to the environmental cue for hydrogenase expression, that is, H2 (Schwartz et al. 1999). At least some of the transcripts initiated at PMBH extend a full 17,000 nt, encompassing hoxA. As a result, PMBH and hoxA form a positive feedback system. Induction of the hydrogenase regulon results in the amplification of both MBH and HoxA. The amplified levels of HoxA also drive up the production of SH. HoxA is, like other members of the NtrC family, a DNA-binding protein (Zimmer et al. 1995). Deletion analysis of the SH upstream region pointed out a tandem palindrome 50 of the 24/ 12 promoter. A similar motif is also present in the MBH upstream region. The distance between the palindromic element and promoter in the two upstream regions is compatible with the standard model for 24/ 12 promoter activation, involving binding of the activator protein and protein-protein contact between activator and sN -RNA polymerase (RNAP) holoenzyme. In vitro studies using HoxAcontaining extracts indicate that HoxA binds to DNA fragments from the SH upstream region harboring the tandem palindrome, suggesting an NtrC-like mechanism of transcriptional activation (Zimmer et al. 1995). Detailed investigation of H2-dependent hydrogenase expression led to the identification of an H2-responsive signaling pathway (Friedrich et al. 1996; Lenz and Friedrich 1998). This pathway is mediated by three components: (1) the response regulator HoxA, (2) a histidine protein kinase designated ‘‘HoxJ,’’ and (3) a cytoplasmic hydrogenase-like protein, called a ‘‘regulatory hydrogenase’’ (RH), encoded by the genes hoxB and hoxC. HoxA and HoxJ constitute a two-component system. As in other two-component systems, HoxJ phosphorylates itself in an autocatalytic reaction. Subsequently, transfer of the phosphate group to the cognate response regulator HoxA can take place. Unlike most other two-component systems, however,

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H2-Metabolizing Prokaryotes

a

PA1

c

PX

PMBH

P1

PA

PB1

P3

P5

UAS hoxKGZMLOQRTV

hypA1B1F1CDEX-hoxABCJ

hupSLCDEFGHIJK

hypABFCDEX

+

+

+ HoxA

J J

PSH

O2

NifA

B B C C

+

+

H2

FnrN

hoxFUYHWI-hypA2B2F2

b

d P2

P3

hupTUV

phupS

hypF

hupSLC

P4 hupDFGHJK

P5

HupR

+

hypAB-hupR-hypCDE

hupSLCDHIR

+ HupR

RegA O2

+ T T V V H2

U

FnrT

+ hynSisp1isp2hynL

U

. Fig. 4.28 Molecular models for the regulation of hydrogenase expression in Ralstonia eutropha (a), Rhodobacter capsulatus (b), Rhizobium leguminosarum (c), and Thiocapsa roseopersicina (d). Colored arrows represent genes or operons. Colored ovals symbolize regulatory proteins. Binding sites of regulatory proteins are indicated by open boxes. See text for details

phosphorylation of the response regulator has a negative effect on transcription of the subordinate genes. This conclusion is based on various experimental findings. First, deletion of gene hoxJ results in a deregulation of the system manifest as a drastic increase in hydrogenase gene expression in the absence of H2. Furthermore, replacement of the conserved Asp-55 residue, the putative phosphorylation target in HoxA, has a similar effect, as does alteration of the Gly-422 in the kinase module of HoxJ (Lenz and Friedrich 1998). HoxB and HoxC comprise a nickelcontaining tetramer with the structure a2b2 (Kleihues et al. 2000; Bernhard et al. 2001). Comparisons of deduced amino acid sequences revealed that HoxBC is a relative of the dimeric [NiFe] hydrogenases. The protein does, indeed, catalyze low but significant rates of H2-dependent methylene blue reduction in vitro (Bernhard et al. 2001). The RH forms a complex with HoxJ (Bernhard et al. 2001). This complex can be reconstituted in vitro by mixing purified components. A stretch of amino acids at the C-terminus of HoxB is essential for both the formation of the a2b2 tetramer and the complex formed between this tetramer and HoxJ (Buhrke 2002). Notably, this C-terminal peptide is different from the C-termini of the energy-generating membrane-bound hydrogenases. In RH null mutants, hydrogenase gene expression is totally abolished, indicating that the RH is

a positive regulator (Lenz and Friedrich 1998). The available data support the following molecular model for H2 sensing (Lenz et al. 2002; > Fig. 4.29): A cytoplasmic H2-sensing complex, consisting of RH and HoxJ, governs the phosphorylation status of HoxA. In the absence of H2, HoxJ mediates phosphotransfer to HoxA, rendering it inactive. When molecules of H2 enter the cell, they engage the RH, unleashing a specific interaction between it and HoxJ. The latter interaction may involve the transfer of electrons between the RH and the PAS domain in the input module of HoxJ. Thus, the RH blocks the net phosphoryl transfer from HoxJ to HoxA and ultimately is a positive control on H2-dependent transcription. Interestingly, the signal transduction pathway is cryptic in some natural isolates of R. eutropha (Lenz et al. 1997). In R. eutropha H16, a single-nucleotide exchange in the region of hoxJ corresponding to the transmitter domain reactivates signal transduction. A second layer of regulation is superimposed on H2responsive signaling. This involves control of hydrogenase expression exerted by additional carbon and energy sources. Catabolite control is probably very important for R. eutropha, since in the natural habitat, it will very likely be confronted with both H2 and organic substrates at the same time. Monitoring

H2-Metabolizing Prokaryotes

H2 Sensor (HoxBC/ HupUV)

Histidine Protein Kinase (HoxJ/HupT)

Response Regulator (HoxA/HupR)

P

a

His Fe Ni

Input

Transmitter

Asp Receiver

Hydrogenase Genes

b H2 H2 H2 Fe Ni

Output

His Input

Transmitter

Asp Receiver

+ Output

. Fig. 4.29 Molecular model for H2 sensing in R. eutropha and R. capsulatus. The upper part of the diagram (a) illustrates the interactions between the components of the H2-sensing apparatus in the absence of H2. The lower part (b) represents the protein–protein interactions in the presence of H2. Domains that are responsible for the biological response of the system under the respective conditions are shown in red. The solid arrow symbolizes the phosphotransfer reaction. Dashed arrows indicate positive (+) or negative ( ) control

hydrogenase gene expression in the presence of both H2 and organic substrates reveals a clear correlation between substrate quality and expression levels: Expression is high when poor substrates are available in addition to H2 and vice versa. In this context, substrate quality is judged on the basis of the corresponding growth rate. The underlying regulatory mechanisms are not yet known. Despite major differences in the organization of the hydrogenase determinants in the phototroph R. capsulatus and the chemolithotroph R. eutropha, the H2-oxidizing systems of these two organisms share a common regulatory mechanism (Vignais et al. 2005). The dimeric [NiFe] hydrogenase of R. capsulatus is encoded in the hupSLC operon under the control of the phupS promoter (Toussaint et al. 1997; > Fig. 4.28b). The hupDFGHJK, hypAB-hupR-hypCDE, and hupTUV operons are transcribed from separate promoters (Elsen et al. 1996). An additional promoter is located upstream of hypF. Promoter phupS is under the control of an NtrC-like regulator, HupR, but remarkably is sD-dependent as evidenced by mutagenesis of promoter sequences (Dischert et al. 1999) and by experiments with rpoN mutants (Colbeau and Vignais 1992). HupR binds to the palindromic sequence TTG-N5-CAA upstream of PhupS (Toussaint et al. 1997; Dischert et al. 1999). An integration host factor (IHF)-binding motif located between the upstream activation sequence (UAS) and phupS is another feature typical of promoters controlled by NtrC-like regulators. As in other systems, the architectural protein IHF is not essential for hydrogenase expression but has a pronounced stimulatory effect

4

(Toussaint et al. 1991). The presence of H2 triggers a 10-fold induction of hydrogenase activity (Toussaint et al. 1997). This effect depends on an H2-sensing system similar to that found in R. eutropha. A histidine protein kinase, HupT, and an H2-sensing hydrogenase, HupUV, cooperate with HupR in mediating signal transduction. HupT is a negative regulator (Elsen et al. 1993; Dischert et al. 1999). HupT autophosphorylates in vitro at the His-217 residue. PhosphoHupT mediates phosphotransfer to the Asp-54 residue of HupR (> Fig. 4.29). As in the case of R. eutropha HoxA, the phosphorylated form of HupR is inactive, preventing transcription of the hydrogenase genes under noninducing conditions (Dischert et al. 1999). HupUV is a cytoplasmically localized dimeric [NiFe] hydrogenase with a low but significant level of hydrogenase activity and catalyzes the H-2H exchange reaction (Vignais et al. 1997, 2000, 2005; Elsen et al. 2003). HupT forms dimers and tetramers in vitro. Under the same conditions, HupUV was present both as a simple heterodimer and in the tetrameric form (HupUV)2. A mixture of the three subunits yielded the complex (HupUV)2-(HupT)2 (Elsen et al. 2003). HupUV presumably interacts with HupT governing the HupT/HupR phosphotransfer reaction. This interaction is, however, diametrically different from that of its R. eutropha counterpart. Since HupUV-mutants express hydrogenase constitutively (Elsen et al. 1996), HupUV must have a stimulatory effect on the net phosphoryl transfer to HupR. Apart from the specific, H2-dependent control, hydrogenase expression is also regulated by the global, redox-responsive regulator pair RegA/RegB (Elsen et al. 2000). RegA exerts a negative effect on hup transcription by binding to two sites overlapping and upstream of phupS. In contrast to R. eutropha and R. capsulatus, hydrogenase expression in the symbiotic N2-fixer R. leguminosarum is not controlled by H2 availability. In the latter organism, hydrogenase is expressed in bacteroids but not in vegetative cells (Palacios et al. 1990). The hydrogenase genes, which belong to at least three transcriptional units, are under the control of O2-sensitive global regulators (> Fig. 4.28c) (Palacios et al. 2005). The first operon, containing the hydrogenase structural genes and an undetermined number of downstream genes, is regulated by NifA (Brito et al. 1997). Transcription is driven by the 24/ 12-type promoter P1 located upstream of hupS (Hidalgo et al. 1992). Deletion analysis of the region 50 of P1 indicated the existence of sequence elements essential for hup gene transcription. Experiments in a heterologous system indicated that transcription from P1 is dependent on sN and is under the control of NifA (Brito et al. 1997). Similar experiments also indicated the involvement of IHF in the activation of P1. Thus, the R. leguminosarum hup genes belong to the nif regulon. The coexpression of hydrogenase and nitrogenase in R. leguminosarum is not surprising, since the physiological role of hydrogenase in this organism is the utilization of H2 generated as a byproduct of N2 fixation. Another promoter designated P3 is located within the hup gene cluster upstream of hupG (Martinez et al. 2004). P3 is sN-dependent and can be mapped to a 24/ 12 sequence element. Like P1, the transcriptional activity of P3 is governed by NifA, but a corresponding UAS is

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missing. P3 directs the transcription of at least the hupGHIJ genes (Martinez et al. 2004). The third transcriptional unit contains the hyp genes with the exception of hypA: hypBFCDEX. Expression of this operon is induced under microaerobic conditions both in bacteroids and in vegetative cells (Palacios et al. 1990). An FNR-dependent promoter, P5, located within hypA controls the expression of the hypBFCDEX operon (Hernando et al. 1995). R. leguminosarum contains two copies of the gene fnrN (Gutierrez et al. 1997). The expression of the hypBFCDEX operon is reduced in single fnrN mutants and abolished in the double mutant. An intriguing finding is the discovery of a defective copy of a gene for a response regulator in the R. leguminosarum hydrogenase gene cluster (Brito et al. 1997). This remnant could be a relic of a defunct H2-dependent regulatory system. The regulation of the multiple hydrogenases of Thiocapsa roseopersicina is complex and not entirely understood. The genome of this bacterium encodes H2-sensing proteins similar to those in R. eutropha and R. capsulatus. However, hydrogenase gene expression in T. roseopersicina is H2-independent. Neither the genes for the putative regulatory hydrogenase (hupUV) nor the gene for a putative histidine kinase (hupT) is transcribed under the experimental conditions tested (Kova´cs et al. 2005a). A response regulator-type protein (HupR) is nevertheless essential for the transcription of the hupSLCDHIR operon (Kova´cs et al. 2005a; > Fig. 4.28d). Accordingly, the hupS gene is preceded by a 24/ 12 region and is RpoN-dependent. In contrast, the expression of the hox1EFUYH operon (encoding a soluble hydrogenase) and of the hynS-isp1isp2-hynLC operon (encoding a second membrane-bound hydrogenase) are not dependent on HupR. The hyn operon is transcribed from a s70 promoter controlled by an FNR homolog (FnrT) (Kova´cs et al. 2005b; > Fig. 4.28d). A basal level of transcription was found in fnrT mutants, suggesting that an additional, FNR-independent promoter is involved in hyn transcription. A very different regulatory mechanism controls expression of the hyc genes which encode the hydrogenase-3 of E. coli. The divergently transcribed hycBCDEFGHI and hypABCDE operons belong to the formate regulon (Bo¨hm et al. 1990; Lutz et al. 1990, 1991). A third transcriptional unit, separated from the hycBCDEFGHI operon by the functionally unrelated genes ascB, -F, and -G, consists of the genes hydN and hypF (Maier et al. 1996b). These operons are transcribed from three sN-dependent promoters designated ‘‘PC,’’ ‘‘PP,’’ and ‘‘P,’’ respectively (> Fig. 4.30). The activator protein FhlA governs the expression of the regulon in response to intracellular formate accumulation (Schlensog and Bock 1990; Schlensog et al. 1994). The FhlA protein forms a complex with formate, binds to sites upstream of the subordinate promoters, and is, itself, transcribed by the weak constitutive, sD-dependent promoter PfhlA. However, transcripts originating at PP encompass fhlA, leading to an amplification of intracellular FhlA levels under inducing conditions. A second regulator, HycA, counteracts FhlA and thus mediates negative control of the regulon. Hydrogenases-1 and 2 of E. coli are encoded by the operons hyaABCDEF and hybOABCDEFG, respectively (Menon et al.

PfhlA fhlA

PP hypABCDE

+ HCOOH

+

PC A

P

hycBCDEFGHI

BF G

NF

+ + FhlA

HycA

. Fig. 4.30 Molecular model for the regulation of the E. coli hyc, hyp, and hyd operons. The genes ascB, ascF, and ascG (green) are not involved in hydrogenase synthesis. See > Fig. 4.28 for details

1990, 1994). Both biochemical and genetic studies have yielded information on the complex regulation of these operons (Ballantine and Boxer 1985, 1986; Sawers et al. 1985; Sawers and Boxer 1986; Brøndsted and Atlung 1994, 1996; Wu et al. 1989; Richard et al. 1999). Like the hyc operon, both the hya and hyb operons are induced under anaerobic conditions. The ArcA/ ArcB system is involved in the regulation of both operons albeit differently. Under anaerobic conditions, ArcA suppresses hyb expression and activates hya expression (Richard et al. 1999). IscR was shown to be a strong repressor of hya expression under aerobic growth conditions and also affects the expression of hybO to some extend (Nesbit et al. 2009). The fumarate and nitrate reduction regulator (FNR) is also involved in the expression of the active holoenzymes, but the effect is evidently indirect. Nitrate, acting via the NarX/NarL and NarQ/NarP systems, represses the synthesis of both hydrogenases. Another twocomponent system, DpiA/DpiB, stimulates expression of the hya operon. The latter effect is mediated by the positive regulator AppY. Expression of the hya operon is largely dependent on the sigma factor sS (Atlung et al. 1997). In an RpoS, mutant expression of hya is reduced in the exponential growth phase, and the strong induction upon entry into stationary phase is abrogated. sS is also involved in the response of hya expression to carbon starvation (Atlung et al. 1997; Pinske et al. 2012). A limitation of ferrous iron due to a mutation in the gene feoB led to a strong decrease in the activity of Hyd-1 and Hyd-2 and to a lesser extent of Hyd-3 (Pinske et al. 2011). However, this effect seems to be primarily posttranslational. The regulation of cyanobacterial hydrogenase genes has been studied in both filamentous and unicellular representatives, and some patterns are emerging (Tamagnini et al. 2007). Under N2-fixing conditions, Gloeothece sp. ATCC 27152 contains an uptake hydrogenase encoded by the hupSLW operon. The latter operon is transcribed only under N2-fixing conditions, that is, in the absence of combined nitrogen (Oliveira et al. 2004). The region immediately upstream of hupS contained a sD-type promoter and a coincident binding motif for the positive transcriptional regulator NtcA. NtcA is an N-responsive regulator which controls N-metabolism in certain cyanobacteria (Herrero et al. 2004). NtcA bound to DNA fragments containing the putative binding site in vitro (Oliveira et al. 2004). A similar scenario has been reported for Lyngbya majuscula CCAP 1446/4

H2-Metabolizing Prokaryotes

(Leitao et al. 2005). In Synechocystis sp. PCC 6803, a bidirectional hydrogenase is encoded by the hoxEFUYH operon. Regulation of this operon is complex involving various parameters including light, availability of combined nitrogen, concentration of Ni in the medium, and anaerobiosis (Oliveira and Lindblad 2009). Transcription is directed by a sD promoter and is dependent on a region between 592 and 690 (relative to the start codon of hoxE) for activation (Gutekunst et al. 2005). A protein bound to a site in this region was identified as LexA. The LexA protein also binds within the region between 198 and 338 bp (Oliveira and Lindblad 2005). Unlike its E. coli homolog, the role of LexA in Synechocystis sp. PCC 6803 is evidently not restricted to the control of DNA repair. It has been postulated that LexA mediates redox-responsive regulation in Synechocystis sp. PCC 6803 (Antal et al. 2006), but this hypothesis is not supported by the data of Kiss and coworkers (Kiss et al. 2009). In addition to LexA, two isologous AbrB-like proteins were shown to bind in the hox upstream region (Oliveira and Lindblad 2008). It is still not clear what physiological factors the above-named regulators respond to and if other regulatory proteins are involved. Cyanobacteria are faced with the problem of reconciling the physiologically inimicable processes of oxygenic photosynthesis and N2 fixation. In heterocystous strains, the nitrogenase is expressed only in specialized cells called heterocysts. Since the physiological role of the cyanobacterial uptake hydrogenase is to recover energy that would otherwise be lost in the form of H2 produced by nitrogenase, it is not surprising that Hup expression is also restricted to the heterocysts. An unusual regulatory mechanism to ensure this sequestration has been reported for the filamentous Anabaena sp. strain PCC 7120 (Carrasco et al. 1995). In the vegetative cells, which do not require uptake hydrogenase, the hupL gene is inactive owing to the presence of a 9.5-kb intervening sequence element. In the course of differentiation to a heterocyst, the intervening element is excised resulting in a continuous hupL reading frame. Excision of the intervening element is the result of a site-specific recombination event targeting 16-bp direct repeats which flank the element. A specific recombinase protein, designated XisC, encoded within the element is necessary and sufficient for its excision (Carrasco et al. 2005). Interrupted hupL genes may be a widespread feature of heterocystous strains, but are not universal (Tamagnini et al. 2000; Happe et al. 2000). An alternative to the spatial sequestration of oxygenic photosynthesis and N2 fixation is temporal separation. The basis for this temporal separation is circadian rhythmicity of gene expression. Schmitz and coworkers showed that in the unicellular cyanobacterium Synechococcus sp. PCC 7942, expression of genes for the bidirectional hydrogenase oscillates in a circadian fashion (Schmitz et al. 2001). Both the hoxEF and hoxUYHW operons are controlled at the level of transcription by a circadian clock. A photoreceptor in the form of bacteriophytochrome is probably responsible for entrainment of the oscillation (Schmitz et al. 2000). Circadian expression of hydrogenase genes has also been demonstrated in other cyanobacteria (Kucho et al. 2005).

4

HrsM

P

P

vhcDGAB

frcADGB

+

+

. Fig. 4.31 Molecular model for the regulation of the selenocysteinecontaining hydrogenases in Methanococcus voltae. The colored arrows represent the vhc and frc operons. Promoters are symbolized by arrows marked ‘‘P.’’ A solid box denotes a negatively acting sequence element. Two positively acting sequence elements are represented by open boxes. Solid triangles denote a repetitive sequence motif

Experimental data on the regulation of hydrogenase genes in archaea are scant. The best known system in this respect is the methanogen Methanococcus voltae (Sorgenfrei et al. 1997). In M. voltae, four operons encode two selenium-containing and two selenium-free hydrogenase isoenzymes (Halboth and Klein 1992). It is, therefore, not surprising that selenium is a key factor governing expression of the hydrogenase genes (Bergho¨fer et al. 1994). The fruADGB and vhuDGAUB operons, which code for the selenium-containing enzymes, are expressed both in the presence and absence of selenium. In contrast, transcripts of the corresponding operons for the selenium-free enzymes are detectable only under selenium limitation. The latter operons, frcADGB and vhcDGAB, are arranged in a head-to-head orientation and are transcribed from divergent promoters located in the common 453-bp upstream region (> Fig. 4.31). Transcription from the frc and fhu promoters is coordinately regulated (Noll et al. 1999). Deletion analysis of the upstream region suggests the presence of both positively and negatively acting sequence elements. One of the negatively acting sequence elements is located between the frc promoter and the start codon of frcA and consists of a tandem repetition of a heptamer (Noll et al. 1999). An undecamer (50 -TCTATATAAAC-30 ) located upstream of each of the promoters was shown to mediate positive control. Interestingly, mutations in either sequence element affect both promoters. A 55-kDa protein which binds specifically to this sequence element has been purified by DNA-affinity chromatography (Muller and Klein 2001). A LysR-type regulator designated HrsM mediates negative control (Sun and Klein 2004). A study on the expression of the M. barkeri ech operon revealed that ech transcript was present under all growth conditions tested, suggesting that ech transcription is constitutive (Ku¨nkel et al. 1998). In M. mazei, the vho and vht operons are differentially expressed (Deppenmeier 1995): vht is expressed during growth on H2 and CO2 but not on acetate. The vho operon, on the contrary, seems to be expressed constitutively. This points to a degree of physiological specialization of the two isoenzymes.

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The anaerobic, hyperthermophilic archaeon Pyrococcus furiosus uses either protons or elemental sulfur as an electron sink to dispose of excess reductant accruing from the fermentation of carbohydrates (Jenney and Adams 2008). In the first case—in the absence of S0—a membrane-bound, multisubunit, Ech-type hydrogenase oxidizes ferredoxin, generating H2 and concomitantly pumps protons across the membrane. The MBH is encoded by the 14-gene operon mbhABCDEFGHIJKLMN (Jenney and Adams 2008). The role of the two cytoplasmic hydrogenases (SHI and SHII) is still unclear. When S0 becomes available, synthesis of all three hydrogenases ceases immediately. At the same time, a membrane-bound enzyme complex encoded by the 13-gene mbx operon and a NAD(P)H-sulfur oxidoreductase (NSR) encoded by the nsr gene are induced (Adams et al. 2001; Schut et al. 2007). Through the concerted action of the latter enzymes, ferredoxin is oxidized, and the electrons are ultimately transferred to S0 yielding H2S. The key regulator controlling this reprogramming of metabolism is SurR (Lipscomb et al. 2009). SurR was identified by massfingerprinting analysis of the proteins bound to the mbh upstream region. A SurR binding motif has been identified by footprinting. Intriguingly, the activity of SurR in in vitro transcription assays was the opposite of what might have been expected on the basis of the expression pattern described above: in vitro SurR activated transcription of the hydrogenase genes and repressed the mbx and nsr genes. SurR apparently functions both as a repressor and as an activator (Lipscomb et al. 2009). The examples discussed above reveal that hydrogenase enzymes are integrated into specific physiological functions by diverse regulatory mechanisms. It is evident that the regulatory scheme is dictated more by the physiological context than by the phylogenetic origin of the given hydrogenase.

[FeFe] Hydrogenases Not much is known about the regulation of genes for [FeFe] hydrogenases. Clostridium acetobutylicum produces high levels of hydrogenase activity during acidogenic fermentation and very low levels after switching to solventogenesis (Gorwa et al. 1996). The monocistronic hydA operon, which encodes an [FeFe] hydrogenase, is expressed as a 1,900-nt transcript (Gorwa et al. 1996). The intracellular levels of hydA mRNA show the same pattern of substrate dependence as the enzyme activity levels, suggesting that hydA is regulated at the level of transcription.

[Fe] Hydrogenases Methanothermobacter marburgensis forms an [Fe] hydrogenase called ‘‘H2-forming methylenetetrahydromethanopterin’’ or ‘‘Hmd.’’ This enzyme is synthesized regardless of the availability of nickel. However, under nickel-limited conditions, the cytoplasmic level of the enzyme is raised sixfold. Under the latter conditions, Hmd substitutes for the F420-reducing hydrogenase,

a [NiFe] hydrogenase, in a reducing step of methanogenesis (> Fig. 4.3). The expression of Hmd is regulated at the transcriptional level and responds to Ni concentration but not to the availability of H2 (Afting et al. 2000). The putative nickel-responsive regulator NikR is believed to upregulate transcription of Hmd and the related of F420-dependent methylenetetrahydromethanopterin dehydrogenase under nickel limitation (Thauer et al. 2010).

Evolutionary Aspects Since the pioneering work of Miller (1953) showing that the synthesis of the simpler organic building blocks of living organisms could occur spontaneously under conditions assumed to be similar to those on the primeval earth, biologists have envisaged an evolution of prebiotic systems in an ocean rich in organic compounds (Miller and Orgel 1974). Hence, it was widely assumed that the first organisms were heterotrophs which fed on the organic compounds in the primeval broth. More recently, discrepancies in this hypothesis have prompted a reconsideration (Maden 1995). One of the incongruities confronting the assumption of a heterotrophic origin of life is the fact that many of the deepest branching lineages of the phylogenetic tree contain autotrophic organisms. This indicates that autotrophy is not of recent origin but rather very ancient. Taking this into account, various hypotheses postulating an autotrophic origin of life have been put forward. The most comprehensive and rigorous of these was proposed by Wa¨chtersha¨user (Wa¨chtersha¨user 1988, 1990, 1992). According to his theory of ‘‘pyrite-pulled surface metabolism,’’ the primordial, energy-yielding process for prebiotic evolution is the reaction of ferrous sulfide and H2S yielding pyrite and H2: FeS þ H2 S ! FeS2 þ H2 The free energy for this reaction under standard conditions is 38 kJ/mol. This is sufficient to drive an archaic CO2-fixing cycle similar to the reductive citric-acid cycle of contemporary organisms. Wa¨chtersha¨user suggests that pyrite formed in this process could serve as a matrix for the growing pool of organic reactants. Anabolic metabolites bind tightly enough to the surface of pyrite preventing their loss to the solution but are still capable of two-dimensional diffusion. The theory outlined above is valuable because of its explanatory power in reconstructing events of prebiotic evolution. It is also attractive because it offers a perspective for later evolutionary phases. Once the formation of phospholipid micelles permitted the liberation of metabolism from the pyrite surface, the above reaction may have been harnessed by early phosphorylation-based energy metabolism. One scenario envisages a primitive system consisting of three enzymes: a sulfur reductase, a hydrogenase, and an ATPase (the latter two being integral membrane proteins). The H2S produced inside the cell by reduction of sulfur could diffuse out and react with FeS yielding pyrite and H2. Thereupon, the H2 could be oxidized by the

H2-Metabolizing Prokaryotes

hydrogenase, providing reductant for the cytoplasmic sulfur reductase. The ensuing H+ gradient could drive ATP synthesis. Wa¨chtersha¨user postulated that a pyrite-based system could catalyze the fixation of CO2 via a mechanism related to the reverse citric acid cycle found in modern microbes. This notion is questionable for thermodynamic reasons (Martin 2012). Various lines of evidence point instead to the Wood-Ljungdahl pathway as the primordial mechanism of CO2 fixation (Russell and Martin 2004; Martin and Russell 2007). Fuchs noted the biochemical antiquity of this pathway, which is the basis of both methanogenic and homoacetogenic metabolism (Fuchs and Stupperich 1985; Fuchs 1986). One of the attractive features of this hypothesis is the fact the Wood-Ljungdahl pathway parallels the inorganic chemistry that is associated with hydrothermal settings (Martin 2012), especially those producing H2-rich exudate in the temperature range 70–100 C. Such niches are characterized by the abiogenic production of reduced carbon compounds including methane, formate and acetate. Thus, the lithoautotrophic metabolism of modern methanogens and homoacetogens preserves the H2/CO2-based chemistry of the first protometabolic systems that evolved in hydrothermal settings. A milestone in the evolution of living systems was the appearance of compartmental organization characteristic of contemporary eukaryotic cells. Many biologists have embraced the notion that the organelles of eukaryotic cells developed out of an endosymbiosis of a bacterium within an archaeal host (Margulis 1970). One of the weak points of this theory is its vagueness regarding the selective pressure responsible for the original endosymbiotic association. Moreover, the classical theory does not explain the origin of hydrogenosomes, the specialized, H2-producing organelles found in many anaerobic lower eukaryotes. A novel hypothesis has been put forward by Martin and Muller (1998) to remedy these deficits. Their ‘‘hydrogen hypothesis’’ proposes that H2 metabolism was the basis of the archaic, endosymbiotic association according to the following scenario: A strictly autotrophic, strictly H2-dependent archaeon such as a primitive methanogen is assumed to be the host cell, and a bacterium with respiratory and fermentative (i.e., H2-producing) capabilities is assumed to be the symbiont. The initial liaison between the two must have originated in an anoxic environment with sufficient H2 and CO2 for growth of the host. The association of symbiont and host would render the host cell independent of environmental sources of H2. On the other hand, in the absence of environmental H2, the host cell would be totally dependent on the symbiont and be subject to selective pressure to optimize gas exchange between the symbiont and itself. This could lead to progressive engulfment of the symbiont. This would, in turn, necessitate that the host cell acquires transport systems (e.g., via gene transfer from the symbiont) in order to supply the symbiont with organic substrates for its heterotrophic metabolism. At this stage, the heterotrophic metabolism of the symbiont and the autotrophic metabolism coexist. In the final stage, a progressive transfer of genetic determinants from symbiont to host could eventually replace its autotrophic metabolism by heterotrophic pathways of the symbiont. This would turn the symbiont into a mitochondrion or a hydrogenosome, depending on the

4

remaining set of enzymes. One of the strengths of the hydrogen hypothesis is the fact that associations like the one postulated as the starting point of the archaeal/bacterial partnership are widespread in present-day microbial communities. The syntrophic associations of H2-producers and H2-consumers in anaerobic habitats are well-known. Another advantage of the hydrogen hypothesis compared with the original endosymbiotic hypothesis is that the former accounts for both mitochondriate and amitochondriate cells. Another aspect of H2 metabolism with a profound impact on the course of evolution is the production of H2 by cyanobacterial mats. With the advent of photosynthesis, the concentration of O2 in the atmosphere began increasing. The gradual transition to an oxidizing atmosphere over a period of about 0.5 billion years set the stage for the evolution of respiratory aerobes. Prerequisite for this development was the production of O2 on a global scale and the concomitant removal of reduced chemical compounds. It was long believed that burial of reduced carbon species was the only process of sufficient magnitude to account for the global change in the atmospheric redox status. Recent studies on subtidal and intertidal mats suggest that H2 production in these mats could have had a major geochemical impact (Hoehler et al. 2001; Jorgensen 2001). The mats, which harbor varied microbial populations dominated by the cyanobacterial species Microcoleus chthonoplastes and Lyngbya spp., were the predominant terrestrial life forms for 2 billion years. Measurements of gas production in the modern cyanobacterial mats revealed a dual cycle of H2 production due in part to the activity of nitrogenase. In contrast to other microbial communities, a major fraction of this H2 is not recycled but liberated into the atmosphere and, hence, eventually escapes into space. Extrapolation of the H2 production by modern mats suggests that, on a geological time scale, they could have contributed significantly to the removal of reductant from the earth’s biogeochemical cycles.

Acknowledgments The authors are indebted to their colleague O. Lenz for helpful comments on the manuscript.

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5 Hydrocarbon-Oxidizing Bacteria Eugene Rosenberg Department of Molecular Microbiology and Biotechnology, Tel Aviv University, Tel Aviv, Israel

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 Effect of Oil Pollution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 202 Isolation and Enumeration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 202 General Nutritional Requirements . . . . . . . . . . . . . . . . . . . . . . 202 Enumeration of Hydrocarbon-Degrading Bacteria . . . . . 204 Enumeration of Hydrocarbon-Degrading Bacteria in Marine Material Not Miscible with Water . . . . . . . 205 Enumeration of Hydrocarbon-Utilizing Bacteria by Direct Plating of Estuarine Water and Sediment Samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 Enumeration of Hydrocarbon-Degrading Bacteria in Freshwater . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 Enumeration of Hydrocarbon-Degrading Bacteria by a 96-Well Plate Procedure . . . . . . . . . . . . . . . . . . . . . . . . 205 Enrichment Culture for Hydrocarbon-Degrading Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 206 Enrichment of Crude Oil-Degrading Bacteria in Supplemented Seawater . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 206 Sequential Enrichment of Hydrocarbon-Degrading Bacteria on Crude Oil in Supplemented Seawater . . . 206 Enrichment of Hydrocarbon-Degrading Bacteria on Bunker C Fuel Oil in Minimal Salts Medium . . . 206 Enrichment of Polyaromatic Hydrocarbon-Degrading Bacteria (PAHs) . . . . . . . . . . 207 Enrichment on Liquid Aromatic Hydrocarbons . . . . 207 Enrichment for Nitrogen-Fixing Hydrocarbon Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 207 Enrichment for Solid Hydrocarbon-Degrading Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208 Identification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208 Physiological Properties . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209 Group-Specific Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209 Physical Interactions Between Bacteria and Hydrocarbons: Adhesion, Desorption, and Emulsification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209

found in many genera, the genera Alcanivorax appear to be special because these bacteria are specialized for growth on hydrocarbons. The initial step in the bacterial degradation of hydrocarbons is the introduction of oxygen into the molecules by group-specific oxygenases. Since these oxygenases are membrane bound, the cell must come into direct contact with their water-insoluble substrate. Hydrocarbon-oxidizing bacteria have potential applications in bioremediation of oil pollution, enhanced oil recovery, production of surface-active agents, and in the use of hydrocarbons as substrates for industrial fermentation processes. 1. 2. 3. 4. 5.

Microbial spoilage of petroleum products Treatment of oil spills and disposal of petroleum wastes Enhanced oil recovery Production of surface-active agents Hydrocarbons as substrates in industrial fermentation processes

Introduction Periodic ecological disasters caused by large oil spills call attention, in a dramatic manner, to the toxicity of petroleum. The fact that hydrocarbons persist for months and even years following major oil spills indicates that hydrocarbon biodegradation is slow in most natural environments. To the microbiologist, the fundamental questions are the following: What are the biochemical mechanisms of hydrocarbon degradation? Which microorganisms are involved? What are their special properties? What limits the rate of hydrocarbon degradation in the environment? And from an applied point of view, what (if anything) can be done to accelerate this rate? Several decades of research on hydrocarbonoxidizing bacteria have provided considerable data relevant to these questions. This chapter will discuss the distribution, nutritional requirements, enumeration, isolation, identification, special physiologic characteristics, and potential applications of hydrocarbon-degrading bacteria. The specific class of methane oxidizers will be presented in a separate chapter.

Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 210

Habitats

Abstract Hydrocarbon-oxidizing bacteria have been isolated from a variety of terrestrial and aquatic environments, using both enrichment and direct plating techniques. Although bacteria able to grow on aliphatic and aromatic hydrocarbons are

Hydrocarbons are a ubiquitous class of natural compounds. Not only are they found in petroleum-polluted areas, but chemical analyses have revealed the presence of significant quantities of aliphatic and aromatic hydrocarbons in most soils and sediments (Giger and Blumer 1974; Stevenson 1966). The most probable origin of the low concentrations of widely distributed

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_66, # Springer-Verlag Berlin Heidelberg 2013

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hydrocarbons is ongoing biosynthesis by certain plants and microorganism (Fehler and Light 1970; Hardwood and Russel 1984; Hunt et al. 1980; Juttner 1976; Kolattukudy et al. 1972; Mikkelson and von Wettstein-Knowles 1978; Winters et al. 1969). Hydrocarbons are produced by reduction of fatty acylCoA by enzymes which utilize NADH or NADPH. Other sources of hydrocarbons are natural seeps on the ocean floor and unburned fuel from oil-burning engines (Floodgate 1984). Since hydrocarbons are natural products as well as pollutants, it is not surprising that hydrocarbon-oxidizing bacteria are widely distributed in nature. A sample of ecological studies on hydrocarbon-degrading bacteria is shown in > Table 5.1. It can be seen that hydrocarbon oxidizers are located in virtually all natural areas, although with large variations in cell concentration. As would be expected, the ratio of hydrocarbonoxidizing bacteria to the total population of heterotrophic bacteria, as well as the variety of hydrocarbon-degrading microorganisms found in a particular ecosystem, may change according to the time of sampling or the extent of oil pollution (Geiselbrecht et al. 1996). Atlas (1981) has discussed many of the factors that limit the growth of hydrocarbon-oxidizing bacteria in nature. These include physical constraints, such as temperature, availability of oxygen, salinity, pH, and the extent to which the particular habitat is an open or closed ecosystem. Nutritional factors are also important and include the availability of utilizable sources of nitrogen, phosphorus, and other elements; the nature of the hydrocarbon substrate and its effective concentration; and the possible presence of toxic substances either in the petroleum product or in the environment itself.

spillage of 55,000 gal of leaded gasoline. The ratio of hydrocarbon-utilizing to total heterotrophic bacteria was reported to be an indicator of the gasoline contamination. These investigators also studied shifts in microbial populations in Arctic coastal water using a continuous flow-through system, following the introduction of an artificial oil slick (Horowitz and Atlas 1977a). The addition of the oil appeared to cause a shift to a greater percentage of petroleum-degrading bacteria. Atlas and Bartha (1973b) found similar results in an oil-polluted area in Raritan Bay off the coast of New Jersey. Hood et al. (1975) compared microbial populations in sediments of a pristine salt marsh with those of an oil-rich marsh in southeastern Louisiana. These investigators also found a high correlation between the percentage of hydrocarbon oxidizers and the level of hydrocarbons in the sediments. Significant increases in the number of hydrocarbon-utilizing microorganisms were found in field soils following the addition of several different oil samples (Raymond et al. 1976). No estimate of the ratio of hydrocarbon oxidizers to the total heterotrophic population was presented. From the studies discussed above, it is clear that the presence of hydrocarbons in the environment frequently brings about a selective enrichment in situ for hydrocarbon-utilizing microorganisms. Evidence also has been presented suggesting that the supplementation of certain ecosystems, particularly oil-polluted marine environments with nitrogen and phosphorus, may increase the relative number of hydrocarbon oxidizers (Atlas and Bartha 1973a; Gutnick and Rosenberg 1977; Reisfeld et al. 1972; Song and Bartha 1990).

Effect of Oil Pollution

The use of hydrocarbons as substrates for bacterial growth presents special problems both to the microorganism using them as a source of carbon and energy and to the investigators in the field of hydrocarbon microbiology. Depending on the solubility of the particular hydrocarbon in water, its physical state (solid, liquid, or gas), and toxicity, different isolation methods must be employed. In all cases, the heterogeneity of the system complicates sampling, enumeration, and growth measurement procedures. After a discussion of general nutritional requirements for hydrocarbon-degrading bacteria, several specific procedures for the selective enrichment and isolation of the different hydrocarbon degraders will be presented.

The localization of hydrocarbon-oxidizing bacteria in natural environments has received considerable attention because of the possibility of utilizing their biodegradation potential in the treatment of oil spills. Because of the enormous quantities of crude and refined oils that are transported over long distances and consumed in large amounts, the hydrocarbons have now become a very important class of potential substrates for microbial oxidation. It is not surprising, therefore, that hydrocarbonoxidizing microorganisms have recently been isolated in large numbers from a wide variety of natural aquatic and terrestrial environments. Several investigators have demonstrated an increase in the number of hydrocarbon-oxidizing bacteria in areas that suffer from oil pollution (> Table 5.1). Walker and Colwell (1976a, b) observed a positive correlation between the percentage of petroleum-degrading bacteria in the total population of heterotrophic microorganisms and the amount of heptane-extractable material in sediments of Colgate Creek, a polluted area of Chesapeake Bay. In contrast, no correlation was found when total numbers of hydrocarbon oxidizers (rather than percentages) were compared to hydrocarbon levels. Horowitz and Atlas (1977b) observed shifts in microbial populations in an Arctic freshwater lake after the accidental

Isolation and Enumeration

General Nutritional Requirements In addition to the requirements for suitable cell-hydrocarbon interactions and the specific genetic potential of the organism for hydrocarbon oxidation, a number of general nutritional conditions must be fulfilled for bacteria to utilize hydrocarbons. These nutritional requirements depend on the fact that hydrocarbons, as the name denotes, are compounds composed solely of carbon and hydrogen atoms. Thus, all other elements essential for cell growth must be available in the growth medium. These

Hydrocarbon-Oxidizing Bacteria

5

. Table 5.1 Sample habitats and characteristics of hydrocarbon-utilizing microorganisms Location

Source

Carbon source

Cell concentration References

Prince William Sound

Surface

Hexadecane

2–12  103 per g

Alaska, beach gravel

Subsurface

Hexadecane

1–12  103 per g

Tyrolean Alps

Subsoil

Diesel

0.2–3  104 per g 3

a

Lindstrom et al. (1991) Margesin and Schinner (1997)

Bayway Refinery, New Jersey Surface

Jet fuel

1–4  10 per g

Surface

Jet fuel

1–6  103 per ga

Prince William Sound

Surface

Fuel oil

103–104 per g

Haines et al. (1996)

Alaska, Duck Island

Sand

–

Puget Sound, Washington

Contaminated sediment

Phenanthrene

104–103 per g

Geiselbrecht et al. (1996)

3

Song and Bartha (1990)

4

Uncontaminated sediment

Phenanthrene

10 –10 per g

Barataria Bay (Louisiana Coast)

Over 200 stations along the coast

Lightweight paraffin oil

103–104 per g mud ZoBell and Prokop (1966)

Chesapeake Bay

Eastern Bay (water)

Nondetergent motor oil

0.5–6  103 per ml Walker and Colwell (1976b)

Eastern Bay (sediment)

Nondetergent motor oil

8–99  103 per g

Walker and Colwell (1976b)

Colgate Creek (water)

Nondetergent motor oil

90–4.4  103 per ml

Walker and Colwell (1976b)

Colgate Creek (sediment)

Nondetergent motor oil

10–9.0  103 per g Walker and Colwell (1976b)

Atlantic Ocean sediment off the North Carolina coast

Alaskan waters

250 m off shore (depth, 9 m) Model petroleum substrate

1.5–1.2  102 per ml

50 km from shore, continental shelf (depth, 62 m)

Model petroleum substrate

3  102 to 3  103 Walker et al. (1976)

375 km from shore (depth, 5,000 m)

Model petroleum substrate

4  104 per ml

Walker et al. (1976)

Chukchi Sea

Crude oil

103–104 per ml

Horowitz and Atlas (1977b)

Port Valdez

Crude oil

3  102 per liter

Robertson et al. (1973)

Prudhoe Bay

Crude oil

7  102 per liter

Atlas and Schofield (1975)

6

Walker and Colwell (1976b)

Cape Simpson oil

Crude oil

3  10 per g soil

Atlas and Schofield (1975)

Airplane Lake

Crude oil

103–102 per g sediment

Crow et al. (1975)

Martigan Point

Crude oil

103–102 per g sediment

Crow et al. (1975)

Marcus Hook, PA

Hexadecane

4.4–11  104 per g Raymond et al. (1976)

Tulsa, OK

Hexadecane

1–5  102 per g

Raymond et al. (1976)

Corpus Christi, TX

Hexadecane

3–66  104 per g

Raymond et al. (1976)

Lake Mendota, WI

Surface water

Hexadecane

102–8  103 per ml Ward and Brock (1976)

Athabasca oil sands

River sediment

Hexadecane

7  104 per ml

Naphthalene

1  103 per ml

Southern Louisiana marsh sediments

Field plots

Wyndham and Costenon (1981)

a

Determined by FDA (fluorescein diacetate) epifluorescence

include molecular oxygen, utilizable forms of nitrogen, phosphorus, sulfur, metals, and trace components. The requirement for molecular oxygen has been given much attention, particularly with respect to maximum production of singlecell protein by hydrocarbon-degrading microorganisms

(Mimura et al. 1973; Schocken and Gibson 1984). The limitation for oxygen is easily overcome in small-scale laboratory studies or in open aqueous systems where the oil–water interface is in direct contact with air at all times. The possibility of anaerobic decomposition of hydrocarbons has received considerable

203

204

5

Hydrocarbon-Oxidizing Bacteria

attention (Hollinger and Zehnder 1996). Although hydrocarbon utilization by strictly anaerobic sulfate-reducing bacteria (e.g., Rosenfeld 1947) has been reported, evidence that pure cultures of sulfate-reducing bacteria can attack hydrocarbon in the absence of additional sources of organic carbon is not definite. However, a few microbial species appear to be able to grow on pure alkane in the absence of molecular oxygen, if provided with nitrate as an electron acceptor (Senez and Azoulay 1961; Mihelcic and Luthy 1988) or sulfate (Rueter et al. 1994; Rabus et al. 1999). The nitrogen and phosphorus requirements for maximum growth of hydrocarbon oxidizers can generally be satisfied by ammonium phosphate. Alternatively, these requirements can be met with a mixture of other salts, such as ammonium sulfate, ammonium nitrate, ammonium chloride, potassium phosphate, sodium phosphate, and calcium phosphate. When ammonium salts of strong acids are used, the pH of the medium generally decreases with growth. This problem can often be overcome by using urea as the nitrogen source. In theory, approximately 150 mg of nitrogen and 30 mg of phosphorus are consumed in the conversion of 1 g of hydrocarbon to cell material. In open systems, the high water solubility of most utilizable sources of nitrogen and phosphorus reduces their effectiveness because of rapid dilution. In principle, this problem can be solved by using oleophilic nitrogen and phosphorus compounds with low C/N and C/P ratios. It was found that a combination of paraffinized urea and octyl phosphate was able to replace nitrate and inorganic phosphate, respectively (Atlas and Bartha 1973a). A more economical way may be to add water-insoluble controlledrelease nitrogen and phosphorus fertilizers. This technology has been successfully demonstrated in laboratory and field experiments (Rosenberg et al. 1996). Another practical source of hydrophobic N and P is guano. One intriguing possibility to obviate the need for addition of nitrogen compounds to the medium is to use a bacterium that is capable of both hydrocarbon degradation and nitrogen fixation. Such microorganisms were reported following enrichment on hydrocarbon media lacking nitrogen salts (Coty 1967). In addition to utilizable sources of nitrogen and phosphorus, the mineral requirements of hydrocarbon-degrading bacteria can be met by the addition of K+, Mg2+, Fe2+, and SO42 to purified media. All other inorganic ions required by bacteria to obtain optimum growth are commonly present in sufficient concentration as contaminants in these salts. For most marine hydrocarbon degraders, artificial seawater (or filtered seawater), supplemented simply with phosphate, a nitrogen source, and the hydrocarbon, serves as an adequate medium for enrichment culture studies. In certain aquatic environments under conditions in which the water was supplemented with nitrogen and phosphorus, a high concentration of iron may limit oil biodegradation (Dibble and Bartha 1976). Under these conditions, an encapsulated oleophilic iron compound, ferric octoate, was found to be as effective in stimulating biodegradation as various water-soluble iron derivatives, such as ferric ammonium citrate.

Enumeration of Hydrocarbon-Degrading Bacteria The determination of the concentration of hydrocarbondegrading bacteria is one of the methods commonly used for monitoring oil pollution in the environment. Theoretical difficulties associated with the interpretation of these data have been discussed elsewhere (Floodgate 1973). The enumeration of hydrocarbon-degrading bacteria presents two special technical problems, sampling and choice of carbon source. Petroleumdegrading bacteria tend to adhere to hydrophobic materials (> Fig. 5.1). Thus, unless the bacteria are removed from the material and dispersed prior to enumeration, only minimum cell numbers can be obtained. The choice of a carbon source is an even more serious problem. Petroleum is an extremely complex mixture of hydrocarbons. Because certain bacteria may grow only on minor components in oil, it would be necessary to incorporate large quantities of petroleum into the growth medium to ensure sufficient substrate for these bacteria to grow well. However, high concentrations of petroleum and mixtures of hydrocarbons cannot be used because they are toxic to bacteria (Vestal et al. 1984). Thus, the enumeration of hydrocarbon-degrading bacteria using petroleum as the carbon source selects primarily for bacteria that can degrade major components of the oil mixture. Often, pure hydrocarbons and mixtures of pure hydrocarbons and fractions of crude oil can be used to advantage in replacing petroleum as the carbon source in the isolation medium. The following four methods have been used to enumerate hydrocarbon-degrading bacteria in the marine, estuarine, and freshwater environments.

. Fig. 5.1 Phase contrast photomicrograph of bacterial strain UP-2 (Horowitz et al. 1975) growing on supplemented 0.1 % hexadecane-seawater medium. During exponential growth, most of the cells appear to be in the form of microcolonies tightly bound to oil droplets. Diameter of oil droplets approximately 150 mm

Hydrocarbon-Oxidizing Bacteria

FeCl3 (25 g · ml 1)

Enumeration of Hydrocarbon-Degrading Bacteria in Marine Material Not Miscible with Water 1. Approximately 2 g of the material to be examined is placed in a sterile bottle containing 100 ml sterile seawater or salts medium (Gunkel and Trekel 1967). 2. After 1 ml of a sterile, nontoxic, nonionic emulsifier and 1 drop of an antifoam agent are added to the sample, the mixture is homogenized to disperse and break up the bacterial aggregates (e.g., an Ultra Turrax homogenizer run at 24,000 rpm for 30 s). 3. The homogenized sample is then diluted serially in steps of 1:10 in sterile seawater or salts medium. 4. One-milliliter samples of the appropriate dilutions are then inoculated into bottles or tubes containing the following sterile medium: Aged seawater

250 ml

NH4Cl

0.5 g

K2HPO4

0.5 g

NaH2PO4

1.0 g

1 drop

Purified agar (Difco)

20 g

Oil powder

10 g

KH2PO4 (10 g · 100 ml 1)a

3.0 ml

K2HPO4 (10 g · 100 ml 1)a

7.0 ml

a

Fungizone

10 mg

a

Added after autoclaving

Enumeration of Hydrocarbon-Degrading Bacteria in Freshwater Basal medium (Ward and Brock 1976): Distilled water

750 g

Distilled water

5

1l

NaCl

0.4 g

NH4Cl

0.5 g

MgSO4 · 7H2O

0.5 g

NaHPO4 · 7H2Oa

0.05 g

KH2PO4a

0.05 g

a

Added after autoclaving

5. After addition of one drop of sterile hydrocarbon, the samples are incubated for 2–6 weeks, depending on the temperature of incubation. 6. Bottles remaining turbid after addition of 1 ml HCl to dissolve inorganic salts are scored, and the most probable number is calculated from tables published in Standard Methods (American Public Health Association 1995).

Serial dilutions are made in the basal medium. One drop of sterile hydrocarbon is added to 10 ml of basal medium. After incubation at the appropriate temperature, growth is detected by pellicle formation at the surface of the oil droplet. Most probable number is determined from the tables published in Standard Methods (American Public Health Association 1995).

Enumeration of Hydrocarbon-Utilizing Bacteria by Direct Plating of Estuarine Water and Sediment Samples

Enumeration of Hydrocarbon-Degrading Bacteria by a 96-Well Plate Procedure

Estuarine salts solution (Colwell et al. 1973): Distilled water

1l

NaCl

10 g

MgCl2

2.3 g

KCl

0.3 g

Oil powder: 10 g of hydrocarbon dissolved in 30 ml of diethyl ether is mixed with 10 g of silica gel, allowed to evaporate, and then added to the following basal medium prior to autoclaving. Oil agar medium: Distilled water

1l

NaCl

10 g

MgSO4

0.5 g

NH4NO3

1.0 g

The 96-well plates were processed with a Beckman Biomek 1000 laboratory robot (Beckman Instruments, Fullerton, CA, USA) which filled the wells with medium, performed tenfold serial dilutions of the sample, and added oil to the inoculated wells (Haines et al. 1996). The robot added 180 ml of BH (BushnellHaas medium, Difco) to each well in 11 of the 12 rows, leaving the first row empty. It transferred 200 ml of undiluted sample to the wells in the first row, mixed their contents, and then transferred 20 ml to each well in the second row. The contents of the second row were mixed, and 20 ml was transferred to each well in the third row. This procedure of mixing and transfer was carried out for all except the last row, which served as a sterile control. Sterile pipet tips were used for each transfer. After the dilutions were completed, 2 ml of oil was added to each well as the growth substrate. The plates were sealed in plastic bags and incubated for 14 days at 20  C. Positive wells were scored in one of two ways. When F2 (number 2 fuel oil) was the carbon source, 50 ml of a sterile solution (3 g · l 1) of INT (iodonitrotetrazolium violet;

205

206

5

Hydrocarbon-Oxidizing Bacteria

Research Organics, Cleveland, OH, USA) was added to each well. INT competes with O2 for electrons from the respiratory electron transport chain, and it is reduced to an insoluble formazan that deposits as a red precipitate in the presence of active respiring microorganisms. Red or pink wells were scored as positive. When a crude oil was used as the carbon source, a smooth oil slick developed in each well. Positive wells were scored by emulsification or dispersion of this oil slick. INT cannot be used effectively with crude-oil substrates because their dark color interferes with detection of formazan deposition.

Enrichment Culture for Hydrocarbon-Degrading Bacteria Since hydrocarbons are natural products that are widely distributed in nature, it is not surprising that bacteria able to degrade hydrocarbons can easily be isolated by standard enrichment culture procedures. By varying parameters, such as temperature, pH, hydrocarbon concentration, and basal medium, a wide variety of different hydrocarbon-degrading and emulsifying bacteria can be obtained from either aquatic or terrestrial environments. In most studies, crude oi1 or a petroleum distillate was used as the sole carbon and energy source in the enrichment culture procedure. Under those conditions, bacteria that specialize in the oxidation of low-molecular-weight n-alkanes are generally obtained. Bacteria that grow more slowly or oxidize minor components of crude oil never increase much in batch enrichments, although the activity of these microorganisms may be of special significance in natural environments. To overcome this difficulty, enrichment culture procedures have to be employed using different carbon sources. The following examples represent only a few of the possible variations.

Enrichment of Crude Oil-Degrading Bacteria in Supplemented Seawater To 20 ml of unsterilized seawater in a 125-ml flask was added 155 mg unsterilized crude oil, 0.056 mM KH2PO4, and 7.6 mM (NH4)2SO4 (Reisfeld et al. 1972). After inoculation with about 1 g beach tar or oily sand, the flask was incubated at 30  C with shaking. After about 1 week, the oil became evenly dispersed throughout the liquid. One milliliter of this culture was then transferred to 20 ml sterile seawater supplemented with 0.056 mM KH2PO4, 7.6 mM (NH4)2SO4, and 1 drop of sterile crude oil. (The crude oil was sterilized by filtration through a Millipore 0.45 mm membrane filter.) After one passage, the oil became emulsified in 2–4 days. Such mixed cultures were maintained by serial transfers to fresh media at 3–4-day intervals. Pure cultures were obtained by streaking the enrichment culture either onto the above medium solidified with 1.5 % agar (Difco) or nutrient agar (Difco) prepared with filtered seawater. Isolated colony types were found to grow both on nutrient- and oil-containing media.

Enrichment culture procedures used to isolate crude oildegrading bacteria, such as that described in the preceding paragraph, yield a mixture of several different strains even after several transfers. One reason for this is the heterogeneity of the carbon source. Low-molecular-weight paraffin oxidizers (C10 to C25) generally dominate the cultures because of their more rapid growth rate. To isolate bacteria that could utilize other fractions of crude oil, the following sequential enrichment culture can be employed:

Sequential Enrichment of Hydrocarbon-Degrading Bacteria on Crude Oil in Supplemented Seawater Inoculate the following sterile medium with a pure culture of an n-paraffin-oxidizing bacterium (which can be obtained by standard enrichment culture procedures): 1 l filtered water, 10 mg K2HPO4 · 3H2O, 450 mg urea, and 0.7 ml crude oil (Horowitz et al. 1975). After 3 days incubation with shaking at 30  C, the residual oil is extracted with 1 l of benzene-pentaneether (3:1:1, v/v/v). The oil remaining after evaporation of the organic solvent in vacuo is referred to as ‘‘bacteria-depleted oil.’’ An enrichment culture is now carried out using the ‘‘bacteriadepleted oil’’ in place of crude oil as the sole source of carbon and energy. The general failure of investigators to isolate microorganism on highly water-insoluble, solid hydrocarbons, such as anthracene, may be due to the fact that the cells remain firmly bound to the substrate. Thus, a standard enrichment culture procedure in which a portion of the bulk water phase is transferred would select against rather than for these specific microorganisms. It may be that for successful enrichments, the solid phase should be used as the inoculum during the sequential transfers.

Enrichment of Hydrocarbon-Degrading Bacteria on Bunker C Fuel Oil in Minimal Salts Medium One gram of beach sand sample or 1 ml of a water sample was added to the following minimal medium containing 0.125 % Bunker C oil (steam-sterilized at 121  C and 15 psi for 15 min in tightly capped flasks to prevent evaporation) (Mulkins-Phillips and Stewart 1974a): Minimal salts medium: Distilled water

1l

NaCl

28.4 g

K2HPO4

4.74 g

KH2PO4

0.56 g

MgSO4

0.50 g

CaCl2

0.1 g

NH4NO3

2.5 g

Trace element stock (pH 7.1)

1 ml

Hydrocarbon-Oxidizing Bacteria

Flasks were incubated at 20  C for 14 days and 120 rpm on a refrigerated gyratory shaker bath. Pure cultures of hydrocarbon-utilizing bacteria were isolated from the enrichment culture by streaking onto minimal salts medium to which 2 % washed Ionagar no. 2 (Oxoid) was added. The carbon source consisted of 0.5 ml of the following hydrocarbon mixture added to sterile filter paper secured in the lids of the petri dishes. The dishes were then inverted and incubated at the appropriate temperature for 1–3 weeks. Hydrocarbon mixture: Naphthalene

0.1 g

Anthracene

0.1 g

Dibenzothiophene

0.1 g

Decalin

5 ml

Hexadecene-1

5 ml

Hexadecane

5 ml

Octadecane

0.1 g

Dodecane

5 ml

Isooctane

5 ml

Enrichment of Polyaromatic Hydrocarbon-Degrading Bacteria (PAHs)

a

C OTTON PLUG

APERTURE VOLATILE HYDROC ARBON

MINERAL SALTS MEDIUM

b

MINERAL SALTS AGAR

Small amounts of fresh sediment known to be contaminated with PAHs were inoculated into the following mineral salts medium (g/l) (Churchill 1999): (NH4)2SO4

10

KH2PO4

5.0

MgSO4 · 7H2O

0.1

Fe(NH)2(SO4)2

0.005

Pyrene

40

Trace metals (Beauchop and Elsden 1960) After adjusting the pH to 7.0 with NaOH, the flasks were shaken for 1 week. Pyrene-degrading bacteria were detected on pyrene-coated mineral medium (as above) agar plates. Zones of clearing around colonies indicated pyrene degradation. The same procedure can be used with other PAHs replacing the pyrene.

Enrichment on Liquid Aromatic Hydrocarbons Liquid aromatic hydrocarbons, such as benzene, toluene, and ethylbenzene, are toxic to bacteria when present in the liquid phase (Gibson 1971). However, if these carbon sources are introduced in the vapor phase, good growth can be obtained. > Figure 5.2 illustrates two methods that can be used for growing bacteria on volatile toxic hydrocarbons. Since the liquid hydrocarbons do not come in direct contact with the salts

5

C OTTON PLUG VOLATILE HYDROC ARBON

. Fig. 5.2 Two methods for the growth of bacteria on volatile hydrocarbons using (a) liquid media and (b) solid media

medium, they need not be sterilized. When the reservoir of volatile hydrocarbons is exhausted, it can easily be refilled with a Pasteur pipette.

Enrichment for Nitrogen-Fixing Hydrocarbon Oxidizers Hydrocarbon-oxidizing bacteria able to grow in the absence of added nitrogen compounds were isolated by addition of 0.1 g soil to 25 ml of mineral salts medium of the following composition (g/l) (Coty 1967): Na2HPO4

0.3

KH2PO4

0.2

MgSO4 · 7H2O

0.1

FeSO4 · 7H2O

0.005

Na2MoO4 · 2H2O

0.002

207

208

5

Hydrocarbon-Oxidizing Bacteria

The containers were incubated in an atmosphere of air and hydrocarbon vapors. After turbidity developed, the cultures were streaked and reincubated on the above mineral salts medium containing 1.5 % washed agar. Purification was achieved after several restreakings and culturing on nitrogenfree mineral salts agar medium. Bacteria able to utilize atmospheric nitrogen on addition of naphthenic acid, n-butane, n-tetradecane, or sodium cyclohexane carboxylate were reported to be isolated by this procedure.

. Table 5.2 Genera of hydrocarbon-degrading bacteria Referencesa Genus

From soil

From aquatic environment

Achromobacter

9, 12, 27

5, 8, 10, 20, 21, 25, 28

Acinetobacter

9, 23, 27, 34

5, 8, 17, 24, 26

Actinomyces

12

4, 28, 31

9, 16, 23, 27

6, 8, 9

Aeromonas

7, 26

Enrichment for Solid Hydrocarbon-Degrading Bacteria

Alcaligenes Arthrobacter

9–12, 14, 15, 18

5, 7, 10, 25

A slurry of soil in 0.1 M phosphate buffer, pH 7.0, with 0.1 % octadecane was incubated for 1 week, with shaking (Miller and Bartha 1989). This enrichment culture was transferred (1:100 ratio) to the following medium (g/l):

Bacillus

14, 23

4, 7, 10, 20, 28

10

5

Na2HPO4

0.4

KH2PO4

0.15

NH4Cl

0.1

MgSO4 · 7H2O

0.02

Iron ammonium citrate

0.005

CaCl2

0.001

Octadecane

0.1

Alcanivorax

Beneckea Brevibacterium

10

Corynebacterium 12, 14, 15, 22, 27 30

Cytophaga

9, 27

Erwinia

1

4

Flavobacterium

1, 13, 15, 22, 27

2, 7, 9, 10, 20, 21 2

Lactobacillus

2

Leucothrix

1

Micrococcus

13, 14, 22 2

Mycobacterium

29

Myxobacterium

3, 14, 23, 32

28

Nocardia

14, 15, 23

2, 4, 5, 10, 21, 26 7

Pseudomonas

1, 9, 11–14, 16, 18, 19, 23, 2, 4, 5, 10, 20, 21, 26, 27, 33 28

Rhodococcus

35

Sarcina

14, 22

Serratia

1

Sphaerotilus

a

2

Moraxella

Peptococcus

Identification

4, 5, 7, 10, 25, 26, 28

Cycloclasticus

Klebsiella

To obtain pure cultures, the enrichment was streaked on the above medium solidified with 2 % agar. The Pseudomonas sp. that was isolated grew on solid alkanes such as hexatriacontane (C36).

The variation in bacterial populations isolated by enrichment culture depends largely on the hydrocarbon substrate used in the enrichment, the culture conditions, and the source of the inoculum. Many species capable of hydrocarbon degradation have been isolated (> Table 5.2). The most frequently isolated bacterial genera are Pseudomonas, Acinetobacter, Flavobacterium, Corynebacterium, Alcanivorax, and Arthrobacter. Most of the investigations on the degradation of aromatic hydrocarbons have been carried out using Pseudomonas putida and species of Beijerinckia and Nocardia (Gibson 1971). Westlake et al. (1974) studied the effect of oil quality and incubation temperature on the genetic composition of hydrocarbon-decomposing populations isolated from an area in British Columbia that had been exposed to chronic pollution with diesel fuel. All of the populations consisted predominantly of Gram-negative rods, including species of Pseudomonas, Acinetobacter, Xanthomonas, Arthrobacter, and Alcaligenes. An extensive study of petroleum-degrading bacteria isolated from Chesapeake Bay waters and sediments was carried out by Austin et al. (1977a, b). A total of 99 strains were examined for

36

20 4

Spirillum

10

20

Vibrio

1, 30

4, 5, 10, 20, 21, 26, 28

Xanthomonas

9, 10

4

Key to references: 1, Atlas et al. (1978); 2, Atlas and Bartha (1972); 3, Antoniewski and Schaefer (1972); 4, Austin et al. (1977a, b); 5, Bartha and Atlas (1977); 6, Bertrand et al. (1976); 7, Buckley et al. (1976); 8, Byrom et al. (1970); 9, Cook et al. (1973); 10, Cundell and Traxler (1973a, b, 1976); 11, Jensen (1975a); 12, Jensen (1975b); 13, Jobson et al. (1972); 14, Jones and Edington (1968); 15, Kincannon (1972); 16, Kiyohara et al. (1982); 17, Makula et al. (1975); 18, McKee et al. (1972); 19, Miller and Bartha (1989); 20, Mironov (1970), Mironov and Lebed (1972); 21, Mulkins-Phillips and Stewart (1974b); 22, Odu (1978); 23, Perry (1977); 24, Reisfeld et al. (1972); 25, Soli (1973); 26, Walker and Colwell (1974), Walker and Colwell (1975), Walker et al. (1976b); 27, Westlake et al. (1974); 28, ZoBell (1964). Adapted from Floodgate (1984) and Bossert and Bartha (1984). 29, Churchill (1999); 30, Geiselbrecht et al. (1996); 31, Barabas et al. (1995); 32, Burback and Perry (1993); 33, Griffol et al. (1994); 34, Ratajczak et al. (1998); 35, Whyte et al. (1998); 36, Cappello and Yakimov (2010)

Hydrocarbon-Oxidizing Bacteria

48 biochemical, cultural, morphological, and physiological characteristics. A statistical analysis revealed 14 phonetic groups, comprising about 85 % of the hydrocarbon-degrading bacteria. These groups were characterized as actinomycetes; coryneforms; Enterobacteriaceae; Klebsiella aerogenes; species of Micrococcus, Nocardia, and Pseudomonas; and Sphaerotilus natans. Special mention should be made of the genus Alcanivorax (Cappello and Yakimov 2010) because these bacteria seem to play a particularly pivotal role in the oil-spill bioremediation (Schneiker et al. 2006). This obligate hydrocarbonoclastic bacterium (petroleum hydrocarbons serve almost exclusively as its source of carbon and energy) is cosmopolitan and found in various marine environments.

Physiological Properties There are two essential characteristics that define hydrocarbonoxidizing bacteria: (1) hydrocarbon-group-specific oxygenases and (2) mechanisms for optimizing contact between the bacterium and the hydrocarbon.

Group-Specific Oxygenases Several reviews have appeared on the microbial metabolism of straight-chain and branched alkanes (Asperger and Kleber 1991; Singer and Finnerty 1984), cyclic alkanes (Perry 1984), and aromatic hydrocarbons (Gibson 1977; Cerniglia 1984; Pe´rezPantoja et al. 2010). It has been established that the first step in the degradation of hydrocarbons by bacteria is the introduction of both atoms of molecular oxygen into the hydrocarbon. In the case of aromatic hydrocarbons, ring fission requires a dihydroxylation reaction, the introduction of two atoms of oxygen, and the subsequent formation of a cis-dihydrodiol (Gibson 1968; Simon et al. 1993). This reaction is catalyzed by a dioxygenase which is a multicomponent, membrane-bound enzyme system (Cerniglia 1992). Further oxidation of the cis-dihydrodiol leads to the formation of catechols that are substrates for another dioxygenase that catalyzes ring fission (Evans et al. 1965). It is important to emphasize that the biochemical mechanism of aromatic hydrocarbon oxidation in prokaryotes is fundamentally different from that of eukaryotes. Fungi and mammalian cells metabolize aromatics using the cytochrome P-450 monooxygenase system, which leads to the formation of arene oxides. These active epoxides can form covalent bonds with nucleophilic sites in DNA, leading to mutations and carcinogenesis. Aromatic hydrocarbons that have been shown to serve as substrates for bacterial oxygenases include benzene, toluene, xylene, naphthalene, phenanthrene, anthracene, benz(a)anthracene, biphenyl, and several of their methylated derivatives. The enzymes necessary for aromatic hydrocarbon degradation are specified, in part, by degradative catabolic plasmids. Enzymes capable of monooxygenating benzene/toluene to phenol/methylphenol and phenols to catechols belong to an

5

evolutionary related family of soluble di-iron monooxygenases (Leahy et al. 2003), which are enzyme complexes consisting of an electron transport system comprising a reductase (and in some cases a ferredoxin), a catalytic effector protein which contains neither organic cofactors nor metal ions and is assumed to play a role in assembly of an active oxygenase (Powlowski et al. 1997), and a terminal hydroxylase with a (abg)2 quaternary structure and a di-iron center contained in each a subunit. These monooxygenases are classified according to their a-subunits, which are assumed to be the site of substrate hydroxylation, into four different phylogenetic groups: the soluble methane monooxygenases, the alkene monooxygenase of Rhodococcus corallinus B-276, the phenol hydroxylases, and the fourcomponent alkene/aromatic monooxygenases (Leahy et al. 2003). In general, alkanes are terminally oxidized to the corresponding alcohol, aldehyde, and fatty acid (Asperger and Kleber 1991). The hydroperoxides may serve as unstable intermediates in the formation of the alcohol (Singer and Finnerty 1984). Fatty acids derived from alkanes are then further oxidized to acetate and propionate (odd-chain alkanes) by inducible b-oxidation systems. The group specificity of the alkane oxygenase system is different in various bacterial species. For example, Pseudomonas putida PpG6 (oct) grows on alkanes of 6–10 carbons in chain length (Nieder and Shapiro 1975), whereas Acinetobacter sp. HOI-N is capable of growth on long-chain alkanes (Singer and Finnerty 1984). The ability of P. putida to grow on C6–C10 alkanes was shown to be plasmid encoded (Chakrabarty et al. 1973). In contrast, all activities necessary for growth of Acinetobacter sp. HOI-N and A. calcoaceticus BD413 appear to be coded by chromosomal genes (Singer and Finnerty 1984). Subterminal alkane oxidation apparently occurs in some bacterial species (Markovetz 1971). This type of oxidation is probably responsible for the formation of long-chain secondary alcohols and ketones. Pirnik (1977) and Perry (1984) have reviewed the microbial oxidation of branched and cyclic alkanes, respectively.

Physical Interactions Between Bacteria and Hydrocarbons: Adhesion, Desorption, and Emulsification The low solubility of hydrocarbons in water, coupled to the fact that the first step in hydrocarbon degradation involves a membrane-bound oxygenase, makes it essential for bacteria to come into direct contact with their hydrocarbon substrates. Two general biological strategies have been suggested for enhancing contact between bacteria and water-insoluble hydrocarbons: (1) specific adhesion mechanisms and (2) emulsification of the hydrocarbon. To understand the special cell-surface properties of bacteria that allow them to grow on hydrocarbons, it is necessary to consider the dynamics of petroleum degradation in natural environments (Rosenberg et al. 1992). Following an oil spill in the sea, the hydrocarbons rise to the surface and come into contact with

209

210

5

Hydrocarbon-Oxidizing Bacteria

air. Some of the low-molecular-weight hydrocarbons volatilize; the remainder are metabolized relatively rapidly by microorganisms, such as Pseudomonas sp., which take up soluble hydrocarbons. These bacteria do not adhere to oil and do not have a high cell-surface hydrophobicity (Rosenberg and Rosenberg 1985). The next stage of degradation involves microorganisms with high cell-surface hydrophobicity, which can adhere to the residual high-molecular-weight hydrocarbons. In the case of A. calcoaceticus RAG-1, this adherence is due to thin hydrophobic fimbriae (Rosenberg et al. 1982). Mutants lacking these fimbriae failed to adhere to hydrocarbons and were unable to grow on hexadecane. Other bacteria exhibit high cell-surface hydrophobicity as a result of a variety of fimbriae and fibrils, outermembrane and other surface proteins and lipids, and certain small cell-surface molecules, such as gramicidin S (Rosenberg et al. 1985) and prodigiosin (Rosenberg et al. 1989). Bacterial capsules and other anionic exopolysaccharides appear to inhibit adhesion to hydrocarbons (Rosenberg et al. 1983). Desorption from the hydrocarbon is a critical part of the growth cycle of petroleum-degrading bacteria. Petroleum is a mixture of thousands of different hydrocarbon molecules. Any particular bacterium is only able to use a part of the petroleum. As the bacteria multiply at the hydrocarbon/water interface of a droplet, the relative amount of nonutilizable hydrocarbon within the droplet continually increases until the cells can no longer grow. For bacteria to continue to multiply, they must be able to move from the depleted droplet to a fresh oil droplet. A. calcoaceticus RAG-1 has an interesting mechanism for desorption and for ensuring that it only reattaches to a droplet of fresh oil. When cells become starved on the ‘‘used’’ hydrocarbon drop or tar ball, they release their capsule. The capsule is composed of an anionic heteropolysaccharide, with fatty acid side chains, referred to as emulsan (Rosenberg 1986). The extracellular, amphipathic emulsan attaches avidly to the hydrocarbon/water interface, thereby displacing the cells to the aqueous phase. Each ‘‘used’’ oil droplet or tar ball is then covered with a monomolecular film of emulsan. The hydrophilic outer surface of the emulsan-coated hydrocarbon prevents reattachment of the RAG-1 cells. The released capsule-deficient bacteria are hydrophobic and readily adhere to fresh hydrocarbon substrate. Many hydrocarbon-degrading microorganisms produce extracellular emulsifying agents (Desai and Banat 1997; Rosenberg and Ron 1997). In some cases, emulsifier production is induced by growth on hydrocarbons (Hisatsuka et al. 1971). Mutants that do not produce the emulsifier grow poorly on hydrocarbons (Itoh and Suzuki 1972). Pretreatment of oil with emulsifying agents can both inhibit and stimulate oil biodegradation (e.g., Foght et al. 1989; Nakahara et al. 1981; Tiehm 1994; Thibault et al. 1996; Liu et al. 1995; Zhang and Miller 1994). As discussed above, emulsification may be a by-product of a cell/ hydrocarbon detachment process. An entire chapter of this book is devoted to bioemulsifiers (Rosenberg and Ron 1997). Acinetobacter sp. HOI-N accumulates extracellular membrane vesicles of 20–50 nm in diameter when grown on

hexadecane (Kappeli and Finnerty 1980). The isolated vesicles partition exogenously supplied hydrocarbons in the form of a microemulsion. These vesicles appear to play a role in the uptake of alkanes. Miller and Bartha (1989) have been able to overcome the difficulties involved in the transport of waterinsoluble, solid hydrocarbons by using unilamellar vesicles. A Pseudomonas isolate grew on octadecane (C18) and hexatriacontane (C36) with Ks values of 2,450 and 2,700 mg·l 1, compared to 60 and 41 mg·l 1, respectively, when the hydrocarbon was presented in the form of liposomes. The data clearly demonstrate the importance of transport in the microbial metabolism of recalcitrant hydrocarbons.

Applications Petroleum microbiology began as an applied subject, and the applied aspects continue to provide the primary impetus for research in this field. Current areas of applied interest are: 1. 2. 3. 4. 5.

Microbial spoilage of petroleum products Treatment of oil spills and disposal of petroleum wastes Enhanced oil recovery Production of surface-active agents Hydrocarbons as substrates in industrial fermentation processes

Biodeterioration of petroleum products, such as fuels, lubricating oils, and oil emulsions, has obvious economic implications. Genner and Hill (1981) have reviewed the data on the microbial spoilage of petroleum products and emphasized that spoilage only occurs when the petroleum products come in contact with water. In addition to avoiding water, spoilage can sometimes be retarded by the use of biocides (Rogers and Kapian 1968) or membrane filtration. In considering the microbial treatment of oil spills, it is essential to distinguish between open systems (e.g., the ocean) and closed ones (e.g., oil storage tanks). In the latter case, it is possible to supplement the system with appropriate sources of nitrogen, phosphates, oxygen, and seed bacteria to enhance microbial growth and petroleum degradation, emulsification, or both. Two early published accounts of the use of these fundamental microbiological principles to enhance oil conversion in a restricted area are the treatment of oily ballast water from an oil tanker (Gutnick and Rosenberg 1977) and of contaminated soil (Raymond et al. 1976). More recently, petroleum pollution has been treated by composting (Kirchmann and Ewnetu 1998), by thermophilic bacteria (Mueller and Nielsen 1996), in soilwater slurries (Zhang and Bouwer 1997), and by using waterinsoluble fertilizers (Rosenberg et al. 1996; Knezevich et al. 2006). In an open system, such as the sea, the ability of resident bacteria to extensively degrade a large oil slick is limited primarily by the concentration of nitrogen and phosphorus. Since there is no economical technology for overcoming these nutrient limitations in an open system, there is at present no practical microbial solution for oil spills at sea.

Hydrocarbon-Oxidizing Bacteria

The use of microorganism in tertiary oil recovery has been the subject of several international conferences and literature reviews (e.g., Westlake 1984; Moses and Springham 1982). After primary and secondary recovery (waterflooding) processes, approximately 70 % of the reservoir oil remains underground, trapped in pore spaces and bound to inorganic minerals. The potential use of microorganisms in situ to release this oil depends on the anaerobic production of organic solvents, such as ethanol and butanol, gasses, such as methane and carbon dioxide, and organic acids. These materials can help overcome the physical forces holding the oil in the reservoir. Also, acid production can dissolve carbonates thus increasing the permeability of the reservoir. In addition, microbial products could enhance oil recovery by producing surface-active material and viscosity-altering polymers. Although the evidence for the positive role of microorganism in enhanced oil recovery is limited to a few poorly controlled experiments (Hitzman 1983), the enormous potential of this technology warrants further investigation. In recent years, interest in bioemulsifiers and other microbial surface-active agents has been growing. Many of these compounds are produced by hydrocarbondegrading microorganisms (Rosenberg 1986; Rosenberg and Ron 1997; Desai and Banat 1997). The advantages of microbially produced surfactants include (1) biodegradability and controlled inactivation, (2) diversity of structure and function for different applications, (3) selectivity for specific hydrocarbon/water interfaces, and (4) characteristic surface modifications. The use of hydrocarbons as inexpensive raw materials for the production of single-cell protein (SCP) was stimulated by the publications of Champagnat and Llewelyn (1962) and Champagnat et al. (1963). During the 1960s, many large oil and fermentation companies were involved in large-scale research and development projects for the conversion of petroleum fractions into SCP. Although the anticipated market for SCP in human and animal nutrition was not realized, these technological developments have provided a rich source of information about how bacteria grow on petroleum, how a continuous process can be scaled-up, and how bulk products can be recovered economically. In the 1970s, several fermentation plants were operating with capacities of 100,000 t of SCP per year. These were the largest biotechnology plants ever built. Because of the increased cost of hydrocarbon feedstock and more stringent governmental regulations governing its use in fermentation industry, there are presently no large-scale commercial fermentation processes based on hydrocarbon substrates. There are, however, a number of excellent microbial processes that have already been developed; these could be activated under the right set of economic conditions. These include processes for producing alcohols, organic acids, and ketones from specific alkanes; single-cell (food) oil from mixed n-paraffins; and large numbers of microbiological metabolites, including vitamins, amino acids, pigments, polysaccharides, enzymes, and alkanes.

5

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6 Lignocellulose-Decomposing Bacteria and Their Enzyme Systems Edward A. Bayer1 . Yuval Shoham2 . Raphael Lamed3 1 Department of Biological Chemistry, The Weizmann Institute of Science, Rehovot, Israel 2 Department of Biotechnology and Food Engineering, Technion – Israel Institute of Technology, Haifa, Israel 3 Department of Molecular Microbiology and Biotechnology, George S. Wise Faculty of Life Sciences, Tel Aviv University, Ramat Aviv, Israel

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 Plant Cell Wall Polysaccharides . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 Cellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 217 Hemicellulose . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Pectin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 221 Cellulose-Degrading Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222 Enzymes That Degrade Plant Cell Wall Polysaccharides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 222 Cellulases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 224 Hemicellulases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Xylan-Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . 225 Mannan- and Galactan-Degrading Enzymes . . . . . . . . 226 Lichenin-Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . 226 Other Polysaccharide-Degrading Activities . . . . . . . . . 226 b D-Xylosidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226 Other Side Chain–Degrading Enzymes . . . . . . . . . . . . . 226 Pectin-Degrading Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . 226 Carbohydrate Esterases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 226 Cellulases and Hemicellulases Are Modular Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 227 The Catalytic Modules: Families of Enzymes . . . . . . . 227 Carbohydrate-Binding Modules (CBMs) . . . . . . . . . . . 228 The Family-9 Cellulases: An Example . . . . . . . . . . . . . . . 230 Cellulase Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 233 Determination of ‘‘True’’ Cellulase Activity: Solubilization of Crystalline Cellulose Substrates . . . 235 Endoglucanase Versus Exoglucanase Activity . . . . . . . 236 Processivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 236 Mechanism of Catalysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 237 Prokaryotic Cellulase Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238 Free Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 238 Multifunctional Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 239 Cellulosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 240 Cell-Bound Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 240 Clostridium thermocellum Cellulosomal Subunits and Their Modules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241 Similarity and Diversity of Scaffoldins from Different Species . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 244

Schematic Comparison of Prokaryotic Cellulase Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245 Free Enzyme Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 245 Multifunctional Enzyme Systems . . . . . . . . . . . . . . . . . . . 247 Cellulosomal Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 248 Gene Regulation of Cellulosomal Components . . . . . . . . . 252 Regulation of Cellulase and Cellulosomal Genes in C. thermocellum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 252 Regulating by Sensing and the Involvement of Alternative Sigma Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 253 New Genetic Tools for C. thermocellum . . . . . . . . . . . . . . . . . 255 Genomics and Metagenomics . . . . . . . . . . . . . . . . . . . . . . . . . . . 255 Phylogenetics of Cellulase and Cellulosomal Systems . . . . . 256 Horizontal Gene Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Gene Duplication . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Domain Shuffling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 256 Proposed Mechanisms for Acquiring Cellulase and Cellulosomal Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 257 Abstract Cellulose and associated polysaccharides, such as xylans, comprise the major portion of the plant cell wall as structural polymers. As the plants evolved and distributed first in the seas and then on land, following their demise, the accumulated cellulosic materials had to be assimilated and returned to nature. Thus the cellulose-degrading bacteria have evolved to complement lignindegrading microbial systems for the purpose of restoring the tremendous quantities of organic components of the plant cell wall to the environment for continued life cycles of carbon and energy on the global scale. This chapter is a sequel to a previous chapter of the same title from the second edition of this treatise (Coughlan MP, Mayer F (1992) The cellulose-decomposing bacteria and their enzyme systems. In: Balows A, Tru¨per HG, Dworkin M, Harder W, Schleifer K-H (eds) The prokaryotes, vol I, 2nd edn. Springer, New York, pp 459–516.) and represents an update of our own subsequent chapter (Bayer EA, Shoham Y, Lamed R (2006) Cellulose-decomposing prokaryotes and their enzyme systems. In: Dworkin M, Falkow S, Rosenberg E, Schleifer K-H, Stackebrandt E (eds) The prokaryotes, vol 2, 3rd edn. Springer, New York, pp 578–617.) which appeared in

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the third edition. Although the basic elements of the previous chapters are still essentially up to date, the field of the cellulosedecomposing bacteria has since advanced greatly, owing to two major factors: (1) the advent, progression, and increasing facility of genome- and metagenome-sequencing efforts and (2) the current initiatives to utilize plant-derived biomass for the production of biofuels as an alternative to fossil fuels for an energy source.

Introduction From an anthropocentric point of view, for millennia, human culture has been intricately involved with cellulose, the major component of the plant cell wall. The development of wood, paper, and textile industries has served to weave cellulosic materials into the fabric of our society. Within the past century, however, cellulosic wastes, derived mainly from the same industries, have also become a major source of environmental pollution. This chapter will concentrate mainly on cellulose and the cellulolytic bacteria, in view of their importance to mankind and world ecology. Nevertheless, the true substrate of these bacteria—i.e., the complement of plant cell wall polysaccharides in general—is much more complex than cellulose alone. Likewise, the complement of enzymes—both the cellulolytic and the non-cellulolytic glycoside hydrolases (GHs)— are produced concurrently in these bacteria for the purpose of efficient synergistic degradation of the complete substrate

composite as it appears in nature. Consequently, when we discuss the cellulose-decomposing bacteria and their enzyme systems, we cannot ignore the related non-cellulolytic enzymes, and these will also be treated, albeit secondarily, in this chapter. The plant cell wall consists of an intricate mixture of polysaccharides (Carpita and Gibeaut 1993); cellulose, hemicellulose, and lignin are its major constituents. These polymers are of a very robust nature. They both equip the plant with a stable structural framework and protect the plant cell from the perils of its environment. Despite its recalcitrant nature, in the guise of dead or dying plant matter, the polysaccharides of the plant cell wall provide an exceptional source of carbon and energy, and a multitude of different microorganisms has evolved which are capable of degrading plant cell wall polysaccharides. In any given ecosystem, the polysaccharide-degrading microbes are not alone, but rely on the complementary contribution of other bacterial and/or fungal species (Bayer and Lamed 1992; Bayer et al. 1994; Ljungdahl and Eriksson 1985). The polymer-degrading strains play a primary and crucial role in the ecosystem by converting the plant cell wall polysaccharides to the respective simple sugars and other degradation products (> Fig. 6.1). They are assisted by satellite microbes, which cleanse the microenvironment from the breakdown products, producing, in the final analysis, methane and carbon dioxide. In a given polysaccharide-degrading microorganism, the enzymes that catalyze the degradation may occur in several possible states, according to recognized paradigms

. Fig. 6.1 Simplified schematic description of a typical ecosystem comprising degrading plant matter. Cellulolytic, xylanolytic, and other hemicellulosic microbes combine to decompose the major polysaccharide components to soluble sugars. ‘‘Satellite’’ microorganisms assimilate the excess sugars and other cellular end products, which are ultimately converted to methane and carbon dioxide

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

(Himmel et al. 2010). These include (1) enzymes in the free state and (2) multifunctional polypeptides in discrete multi-enzyme complexes and/or enzymes attached directly to the bacterial cell surface. All of these paradigms exhibit similar types of enzyme components, which usually comprise modular proteins that contain a multiplicity of functional modules. The ‘‘free’’ enzymes comprise a single polypeptide chain, which contains a catalytic module usually connected to a cellulose-binding module (CBM). The multifunctional enzyme paradigm includes more than one catalytic module per polypeptide chain. Cellulosomes are exocellular macromolecular machines, designed for efficient degradation of cellulose and associated plant cell wall polysaccharides (Bayer et al. 1998a, 2004, 2008; Shoham et al. 1999; Doi and Kosugi 2004). In contrast to the free and multifunctional enzymes, the cellulosome complex is composed of a collection of subunits, each of which comprises a set of interacting functional modules. Thus, one type of cellulosomal module, the CBM, is selective for binding to the substrate. Another family of modules, the catalytic modules, is specialized for the hydrolysis of the cellulose chains. Yet another complementary pair of modules—the cohesins and dockerins— serves to integrate the enzymatic subunits into the complex and the complex, in turn, into the cell surface. Multiple copies of the cohesins form a unique type of nonenzymatic integrating subunit called scaffoldin to which the dockerin-containing enzymes are attached. This ‘‘Lego™’’-like arrangement of the modular subunits generates an intricate multicomponent complex, the enzymes of which are bound en bloc to the insoluble substrate and act synergistically toward its complete digestion. Finally, single enzymes and cellulosomes can both be attached to the bacterial cell surface using one of several mechanisms. In the case of a single enzyme, the catalytic module is attached to the cell surface via a specialized binding module. In the cellulosome, a similar type of binding can occur, but this is done so via an anchoring scaffoldin which then binds to the primary (enzymeintegrating) scaffoldin. Alternatively, in some cases, the enzyme or anchoring scaffoldin is bound covalently to the cell surface enzymatically. Finally, the attachment of some types of cellulosome to the bacterial surface has not yet been elucidated. The above paradigms will be discussed in more detail later in this chapter. A list of cellulose-degrading bacteria is presented in > Table 6.1, together with their dominant enzyme paradigm (some species show numerous types of enzymes, e.g., free and cellulosomal) and their distinctive types of modules (i.e., CBMs, cohesins, dockerins). Inherent to the study of cellulases and related enzymes is their potential industrial application—particularly toward conversion of cellulosic biomass to biofuels. For reviews on the potential uses of these enzymes, the reader is referred to appropriate reviews on the subject (Bhat 2000; Himmel et al. 1999, 2007; Lynd et al. 1991, 2008; Perlack et al. 2005; Ragauskas et al. 2006; Schubert 2006; Galbe and Zacchi 2007; DOE 2008; Himmel 2008; Wall et al. 2008; Nordon et al. 2009; Sheehan 2009; Wilson 2009; Xu et al. 2009; Klein-Marcuschamer et al. 2011).

6

Plant Cell Wall Polysaccharides Plant cells produce a composite matrix of hardy and durable polysaccharides on the outer surface of the plasma membranes, called the cell wall (Carpita and Gibeaut 1993). The cell wall confers a protective coating to the plant cell, providing structure, turgidity, and durability, which render the cell resistant to the outer elements, including mechanical, chemical, and microbial assault. Different types of plant cell tissues exhibit different ratios of the three major types of cell wall component; on the average, the cell wall contains roughly 40 % cellulose, 30 % hemicellulose, and 20 % lignin, but the exact composition of an individual type of plant varies greatly. The first two polymers are indeed polysaccharides. On the other hand, lignin is a heterogeneous, high-molecular-weight hydrophobic polymer, which consists of non-repeating aromatic monomers connected via phenoxy linkages (Higuchi 1990; Lewis and Yamamoto 1990). Unlike cellulose and hemicellulose, which are degraded aerobically or anerobically, lignin degradation requires oxygen and is limited to filamentous prokaryotes (e.g., the Actinomycetes Streptomyces viridans) and fungi (e.g., Phanerochaete chrysosporium, Bejerkendera adusta, and Pleurotus ostreatus), which produce a complicated set of enzymes that hydrolyze the polymer. In fact, the recalcitrant lignin interferes severely with the access of enzymes to the cellulose component, and is rate-limiting for anaerobic degradation of cellulose. In any case, the lignin component must be degraded or removed, before efficient degradation of cellulose can take place. Nevertheless, since lignin is not a polysaccharide, it will not be discussed further in this chapter.

Cellulose Cellulose is the major constituent of plant matter and thus represents the most abundant organic polymer on Earth. Cellulose is a remarkably stable homopolymer, consisting of a linear (unbranched) polymer of b-1,4-linked glucose units. Chemically, the repeating unit is simply glucose, but, structurally, the repeating unit is the disaccharide cellobiose, i.e., 4-O-(b-Dglucopyranosyl)-D-glucopyranose, since each glucose residue is rotated 180 relative to its neighbor (> Fig. 6.2). The individual cellulose chains contain from about 100 to more than 10,000 glucose units, packed tightly in parallel fashion into microfibrils by extensive inter and intrachain hydrogen bonding interactions, which account for the rigid structural stability of cellulose. The microfibrils exhibit variable amounts of crystalline and amorphous components, again depending on the degree of polymerization, the extent of hydrogen bonding and, ultimately, on the source of the cellulose. Cellulose of the plant cell wall is composed of two different forms: cellulose Ia and cellulose Ib. Cellulose Ia is in a triclinic state with a single chain per unit cell and is of higher energy than cellulose Ib, which is in a monoclinic state and much more stable (Atalla and VanderHart 1984; Sugiyama et al. 1991; Atalla 1999; Ding and Himmel 2006). Enzymatic hydrolysis of the Ia form occurs more readily, but the cellulose of the plant cell wall comprises mainly

217

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Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Table 6.1 A list of cellulose-degrading bacteria Bacterium

Distinctive enzyme paradigm

Distinctive modular components

Phylum; Class

‘‘Acetivibrio cellulolyticus’’ CD2a

Cellulosomal

Cohesins, Dockerins and CBM3s

(Firmicutes; Clostridia)b

Acidothermus cellulolyticus 11B ATCC 43068

Free

CBM2s and CBM3s (often together in the Actinobacteria; same protein) Actinobacteria

Actinoplanes sp. SE50/110

Free

CBM2s

Actinobacteria; Actinobacteria

Actinosynnema mirum DSM 43827

Free

CBM2s

Actinobacteria; Actinobacteria

Amycolatopsis mediterranei S699 and U32

Free

CBM2s

Actinobacteria; Actinobacteria

Anaerocellum thermophilum DSM 6725

Multi-functional

CBM2s

Firmicutes; Clostridia

Bacillus cellulosilyticus DSM 2522

CBM5s

Firmicutes; Bacilli

Bacillus licheniformis ATCC 14580/ DSM13

—

Firmicutes; Bacilli

—

Firmicutes; Bacilli

Cohesins, Dockerins

(Firmicutes; Clostridia)b

Bacillus pumilus SAFR-032 ‘‘Bacteroides cellulosolvens’’ ATCC 35603a

Cellulosomal

Butyrivibrio fibrisolvens 16/4

Free

Caldicellulosiruptor kronotskyensis 2002 Multi-functional

CBM2

Firmicutes; Clostridia

CBM3s

Firmicutes; Clostridia

Caldicellulosiruptor lactoaceticus 6A

Multi-functional

CBM3s

Firmicutes; Clostridia

Caldicellulosiruptor obsidiansis OB47

Multi-functional

CBM3s

Firmicutes; Clostridia

Caldicellulosiruptor saccharolyticus DSM Multi-functional 8903

CBM3s

Firmicutes; Clostridia

Catenulispora acidiphila DSM 44928

Free

CBM2s

Actinobacteria; Actinobacteria

Cellulomonas fimi ATCC 484

Free

CBM2s

Actinobacteria; Actinobacteria

Cellulomonas flavigena DSM 20109

Free

CBM2s

Actinobacteria; Actinobacteria

Cellulosilyticum ruminicolaa

Free

CBM3s

Firmicutes; Clostridia

Cellvibrio gilvus ATCC 13127

Free

CBM2s

Actinobacteria; Actinobacteria

Cellvibrio japonicus Ueda107

Free

CBM2s, CBM5s

Actinobacteria; Actinobacteria

Clostridium acetobutylicumc ATCC 824 and EA 2018

Cellulosomal

Cohesins, Dockerins

Firmicutes; Clostridia

Clostridium cellulolyticum H10 ATCC 35319

Cellulosomal

Cohesins, Dockerins

Firmicutes; Clostridia

Clostridium cellulovorans 743B

Cellulosomal

Cohesins, Dockerins

Firmicutes; Clostridia

Clostridium clariflavum DSM 19732

Cellulosomal

Cohesins, Dockerins and CBM3s

Firmicutes; Clostridia

Clostridium josuia

Cellulosomal

Cohesins, Dockerins

Firmicutes; Clostridia

Clostridium lentocellum DSM 5427

Free

CBM2s and CBM3s

Firmicutes; Clostridia

Clostridium phytofermentans ISDg

Free

CBM3

Firmicutes; Clostridia

Clostridium sp. BNL1100

Cellulosomal

Cohesins, Dockerins

Firmicutes; Clostridia

Clostridium stercorarium

a

Clostridium thermocellum ATCC 27405 and DSM 1313

Free

CBM3s

Firmicutes; Clostridia

Cellulosomal

Cohesins, Dockerins and CBM3s

Firmicutes; Clostridia

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Table 6.1 (continued) Distinctive enzyme paradigm

Bacterium a

Distinctive modular components

Phylum; Class

Free

CBM3

Firmicutes; Clostridia

Fibrobacter succinogenes subsp. succinogenes S85

Cell surface

CBM6s and CBM35s

Fibrobacteres/ Acidobacteria group

Hahella chejuensis KCTC 2396

Free

CBM2s

Proteobacteria; Gammaproteobacteria

Eubacterium cellulosolvens

Herpetosiphon aurantiacus ATCC 23779

Free

CBM2s

Chloroflexi; Chloroflexi

Jonesia denitrificans DSM 20603

Free

CBM2s

Actinobacteria; Actinobacteria

Mahella australiensis 50-1 BON

Free

CBM64s

Firmicutes; Clostridia

Micromonospora aurantiaca ATCC 27029 Free

CBM2s

Actinobacteria; Actinobacteria

Micromonospora sp. L5

Free

CBM2s

Actinobacteria; Actinobacteria

Nocardiopsis dassonvillei subsp. dassonvillei DSM 43111

Free

CBM2s

Actinobacteria; Actinobacteria

Paenibacillus barcinonensisa

Free

CBM3s

Firmicutes; Bacilli

Paenibacillus mucilaginosus KNP414

Free

CBM3s

Firmicutes; Bacilli

Paenibacillus polymyxa E681, M1 and SC2

Free

CBM3s

Firmicutes; Bacilli

Paenibacillus terrae HPL-003

Free

CBM3s

Firmicutes; Bacilli

Ruminococcus albus 7 (8, 20 and F40)

Cell surface

CBM37s and Dockerins (Single cohesin)

Firmicutes; Clostridia

Ruminococcus sp. 18P13 a

Cellulosomal?

Cohesins, Dockerins

Firmicutes; Clostridia

a

Cellulosomal

Cohesins and Dockerins

Firmicutes; Clostridia

Saccharophagus degradans 2-40

Free

CBM2s and CBM10s

Proteobacteria; Gammaproteobacteria

Salinispora arenicola CNS-205

Free

CBM2s

Actinobacteria; Actinobacteria

Salinispora tropica CNB-440

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces avermitilis MA-4680

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces bingchenggensis BCW-1

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces coelicolor A3 (2)

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces flavogriseus ATCC 33331

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces hygroscopicus subsp. jinggangensis 5008

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces scabiei 87.22

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces sp. SirexAA-E

Free

CBM2s

Actinobacteria; Actinobacteria

Streptomyces venezuelae ATCC 10712

Free

CBM2s

Actinobacteria; Actinobacteria

Streptosporangium roseum DSM 43021

Free

CBM2s

Actinobacteria; Actinobacteria

Ruminococcus flavefaciens

6

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Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Table 6.1 (continued) Bacterium

Distinctive enzyme paradigm

Distinctive modular components

Phylum; Class

Teredinibacter turnerae T7901

Free

CBM2s and CBM10s

Proteobacteria; Gammaproteobacteria

Thermobifida fusca YX

Free

CBM2s

Actinobacteria; Actinobacteria

Thermobispora bispora DSM 43833

Free

CBM2s

Actinobacteria; Actinobacteria

Verrucosispora maris AB-18-032

Free

CBM2s

Actinobacteria; Actinobacteria

Xylanimonas cellulosilytica DSM 15894

Free

CBM2s

Actinobacteria; Actinobacteria

a

The genome sequences for these species have not yet been released (March, 2012) Lin et al. (1994) c C. acetobutylicum is not considered to be a cellulolytic bacterium. However, for the purposes of the present chapter, the presence of a cellulosome gene cluster, containing a multiple cohesin-bearing scaffoldin and dockerin-containing enzymes (including a GH48 enzyme), justifies, in view of the authors, its inclusion in the table b

. Fig. 6.2 Structure of cellulose. Three parallel chains are shown, and a glucose moiety and repeating cellobiose unit are indicated. The model was built by Dr. Jose´ Tormo, based on early crystallographic data

the more stable Ib form. The hydroxyl groups of glucose are in the equatorial position, as opposed to the axial positions which are all nonpolar protons that do not participate in hydrogen bonding interactions. Thus, owing to the packing of the glucose chains in the microfibrils, the ‘‘sides’’ are polar and hydrogen bonding whereas the ‘‘tops and bottoms’’ are hydrophobic in

character (Matthews et al. 2006). The microfibrils themselves are further assembled into plant cell walls, the tunic of some sea animals, pellicles from bacterial origin, etc. Highly crystalline forms of cellulose include cotton, bacterial cellulose (from Acetobacter xylinum) and the cellulose from the algae Valonia ventricosa, which exhibit crystallinity levels of about 45 %, 75 %,

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

and 95 %, respectively. The following reviews are available for more information on the structure of cellulose: A talla (1999); Atalla and VanderHart (1984); Chanzy (1990); O’Sullivan (1997); Ding and Himmel (2006); Moon et al. (2011).

Hemicellulose Hemicelluloses are relatively low-molecular-weight, branched heteropolysaccharides that are associated with both cellulose and lignin and together build the plant cell wall material (Puls and Schuseil 1993; Timell 1967). The main backbone of hemicellulose is usually made of one or two sugars, which determines their classification. For example, the main backbone of xylan is composed of 1,4-linked-b-D-xylopyranose units. Similarly, the backbone of galactoglucomannan is made of linear 1,4-linked bD-glucopyranose and b-D-mannopyranose units with a-1,6linked galactose residues. Other common hemicelluloses include arabinogalactan, lichenins (mixed 1,3-1,4-linked b-D-glucans), and glucomannan. Most hemicellulases are based on a 1,4-blinkage and the main backbone is branched, whereas the individual sugars may be acetylated or methylated. For example, the linear xylan backbone is highly substituted with a variety of saccharide and nonsaccharide components (> Fig. 6.3). In the plant cell wall, xylan is closely associated with other wall

6

components. The 4-O-methyl-a-D-glucuronic acid residues can be ester-linked to the hydroxyl groups of lignin, providing cross-links between the cell walls and lignin (Das et al. 1984). Similarly, feruloyl substituents serve as cross-linking sites to either lignin or other xylan molecules. Thus, the chemical complexity of xylan is in direct contrast to the chemical simplicity of cellulose. Likewise, the structural diversity of the xylans is in contrast to the structural integrity of the cellulose microfibril. Consequently, unlike the crystalline-like character of cellulose, the hemicellulose component adopts a gel-like consistency, providing an amorphous matrix in which the rigid crystalline cellulose microfibrils are embedded.

Pectin Pectin is a structural polysaccharide which is another major component of the primary cell wall of terrestrial plants. Pectin derivatives serve to mediate plant defense responses and regulate plant development (Ridley et al. 2001). The pectins are heteropolysaccharides composed of a-(1-4)-linked galacturonic acid, substituted with numerous constituent groups, e.g., xylose, rhamnose, and galactose. In addition, a large percentage of the galacturonic carboxyl groups are methylated. During the normal physiological processes of the plant (including plant

. Fig. 6.3 Composition of a typical xylan component of hemicellulose. The xylobiose unit (b-Xylp–b-Xylp) is indicated by the blue-sided box, as are major substituents: Me-a-GlcA, methylglucuronic acid; aAraf, arabinofuranosyl group; OAc, acetyl group. A presumed lignin attachment site to a feruloyl substituent of xylan is also illustrated. Sites of cleavage by selected hemicellulases and carbohydrate esterases are also shown: Xyn, xylanase; Abn, arabinofuranosidase; Glr, glucuronidase; Axe, acetyl xylan esterase; Fae, ferulic acid esterase

221

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Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

growth, maturation, fruit ripening, and aging), the distribution, quantity, chemical composition, and structure of pectin is altered.

Cellulose-Degrading Bacteria The cellulolytic microbes occupy a broad range of habitats. Some are free living and rid the environment of plant polysaccharides by converting them to the simple sugars which they assimilate. Others are linked closely with cellulolytic animals, residing in the digestive tracts of ruminants and other grazers or in the guts of wood-degrading termites and worms (Haigler and Weimer 1991). Cellulose-based ecosystems include soils, swamps, marshes, rivers, lakes, and seawater sediments; rotting grasses, leaves, and wood; cotton bales; sewage sludge; silage; compost heaps; muds; and decaying vegetable matter in hot and volcanic springs, acid springs, and alkaline springs (Ljungdahl and Eriksson 1985; Stutzenberger 1990). The cellulolytic microorganisms include protozoa, fungi, and bacteria and are ubiquitous in nature. The cellulosedecomposing bacteria include aerobic, anaerobic, mesophilic, and thermophilic strains, inhabiting a great variety of environments, including the most extreme, vis-a`-vis temperature, pressure, and pH. Cellulolytic bacteria have also been found in the gut of wood-eating worms, termites, and vertebrate herbivores, all of which exploit anaerobic symbionts for the digestion of wood and fodder. In nature, many cellulolytic species exist in symbiotic relationships with secondary microorganisms (Ljungdahl and Eriksson 1985). The primary microorganisms degrade cellulose directly to cellobiose and glucose. Only part of the breakdown products is assimilated by the polymer degrading strain(s), and the rest is utilized by the satellite microorganisms. Removal of the excess of sugars promotes further cellulose degradation by the primary species, since cellobiose-induced inhibition of cellulase action and repression of cellulase synthesis are precluded. Modern interest in cellulolytic microorganisms was spawned by the decay of cotton fabric in army tents and military clothing in the South Pacific jungles during World War II. The basic research program that resulted from this military problem led to the establishment of the US Army Natick Laboratories (Reese 1976). The resultant research led to the discovery that the causative agent for the costly problem was a cellulolytic fungi, Trichoderma viride (subsequently renamed Trichoderma reesei). Subsequent research, originally from the Natick Laboratories and later spreading to other research institutes and universities, led to the identification and classification of thousands of different strains of cellulolytic fungi and bacteria. Many of the major types of cellulolytic bacteria have been listed in the chapter published in the second edition of The Prokaryotes (Coughlan and Mayer 1992). During the interim period until publication of the chapter in the third edition (Bayer et al. 2006), the major emphasis in the area did not concentrate on the discovery or description of new cellulolytic strains but centered on

characterizing the enzymes and enzyme systems from selected bacteria that degrade cellulose and plant cell wall polysaccharides in general. More recently, however, the emerging simplicity of genome sequencing efforts and metagenomic prospecting (Li et al. 2009) has supplanted the more tedious biochemical approaches.

Enzymes That Degrade Plant Cell Wall Polysaccharides The chemical and structural intricacy of plant cell wall polysaccharides is matched by the diversity and complexity of the enzymes that degrade them. The cellulases and hemicellulases are family members of the broad superfamily of glycoside hydrolases (see > Table 6.2), which catalyze the hydrolysis of oligosaccharides and polysaccharides (Gilbert and Hazlewood 1993; Kuhad et al. 1997; Ohmiya et al. 1997; Schu¨lein 1997; Tomme et al. 1995a; Viikari and Teeri 1997; Warren 1996; Wilson and Irwin 1999). In the past decade, numerous bacterial genomes were sequenced (see > Table 6.1), and databases for the rapidly spiraling accumulation sequences and structures of cellulolytic and hemicellulolytic enzymes are readily available online (see discussion below). Historically, the type of substrate and manner in which a given enzyme interacts with its substrate were decisive in the classification of the glycosidases, as established first by the Enzyme Commission (EC) and later by the Nomenclature Committee of the International Union of Biochemistry. Enzymes were usually named and grouped according to the reactions they catalyzed. Thus, cellulases, xylanases, mannanases, and chitinases were grouped a priori in different categories. Moreover, enzymes which cleave polysaccharide substrates in the middle of the chain (‘‘endo’’-acting enzymes) versus those which clip at the chain ends (‘‘exo’’-acting enzymes) were also placed in different groups. For example, in the case of cellulases, the endoglucanases were grouped in EC 3.2.1.4, whereas the exoglucanases (i.e., cellobiohydrolases) were classified as EC 3.2.1.91. It is interesting that the distinction between endo- and exoacting enzymes is also reflected by the architecture of the respective class of active site, even within the same family of enzymes (> Fig. 6.4). The endoglucanases, e.g., are commonly characterized by a groove or cleft, into which any part of a linear cellulose chain can fit. On the other hand, the exoglucanases bear tunnellike active sites, which can only accept a substrate chain via its terminus. The exo-acting enzyme apparently threads the cellulose chain through the tunnel, wherein successive units (e.g., cellobiose) would be cleaved in a sequential manner. The sequential hydrolysis of a cellulose chain is a relatively new notion of growing importance, which has earned the term ‘‘processivity’’ (Davies and Henrissat 1995), and processive enzymes are considered to be key components which contribute to the overall efficiency of a given cellulase system. Though instructive, there is growing dissatisfaction with the endo/exo terminology. As our understanding of the nature of

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Table 6.2 Major glycoside hydrolase families involved in the degradation of plant cell wall polysaccharides and their enzymatic activities. The glycoside hydrolase families (GHn) in which some members exhibit standard cellulase activities are shown in bold (See CAZy Website for more details: http://www.cazy.org/) GH family GH1

GH2

GH3

GH5

GH6

Enzymes

Catalytic mechanism

Numerous activities, including bglucosidase, b-galactosidase, bmannosidase, and b-glucuronidase; but not b-xylosidase activity

Retaining

Numerous activities, including bgalactosidase, b-mannosidase, and bglucuronidase; but neither b-glucosidase nor b-xylosidase activities

Retaining

Numerous activities, notably bRetaining glucosidase and b-xylosidase activities; but also glucan 1,3-b-glucosidase, glucan 1,4-b-glucosidase and exo-1,3(4)glucanase activities Retaining Broad spectrum of cellulase and hemicellulase activities, including endoglucanase, xylanase, 1,3-bmannanase; b-mannosidase, glucan 1,3-bglucosidase, licheninase, glucan endo-1,6b glucosidase, mannan endo-1,4-bmannosidase, endo-1,6-b-galactanase, and xyloglucan-specific endo-1,4-bglucanase activities Cellulase activities in both aerobic bacteria Inverting and fungi (not found in archaea): both endo- and exo-glucanase (cellobiohydrolase) activities

GH7

Cellulase activities exclusive to the fungi: both endo- and exo-glucanase (cellobiohydrolase) activities

Retaining

GH8

Major cellulase family, with additional members that exhibit lichenanase and xylanase activities

Inverting

GH9

Inverting Endo-, processive endo-, and exoglucanase (cellobiohydrolase) activities in bacteria, plants, and fungi (but rare in archaea)

. Table 6.2 (continued) GH family

Enzymes

Catalytic mechanism

GH19

Chitinases

Inverting

GH26

b-Mannanase and 1,3-b-xylanase activities Retaining

GH30

1,6-b-Glucanase and b-xylosidase activities

GH39

b-Xylosidase activity

Retaining

GH42

b-Galactosidase activity

Retaining

GH43

Broad spectrum of hemicellulase activities, Inverting including xylanase, arabinanase, barabinofuranosidase, b-xylosidase, and galactan 1,3-b-galactosidase activities in bacteria and fungi

GH44

Endoglucanase and xyloglucanase activities, mainly in bacteria

Retaining

GH45

Endoglucanase activity, mainly in fungi (some bacteria)

Inverting

GH47

a-Mannosidase activity, mainly in fungi

Inverting

GH48

Cellobiohydrolases acting from the reducing ends of cellulose and endoprocessive cellulases; mainly in bacteria; an important enzyme in all cellulosomes and in some non-cellulosomal systems

Inverting

GH51

a-L-Arabinofuranosidase and endoglucanase activities

Retaining

GH52

b-Xylosidase activity

Retaining

GH53

Endo-1,4-b-galactanase activity

Retaining

GH54

a-L-Arabinofuranosidase and b-xylosidase Retaining activities, mainly in fungi

GH55

Exo- and endo-1,3-glucanase activities, mainly in fungi

Inverting

GH62

a-L-Arabinofuranosidase activity

Unknown

GH64

1,3-b-Glucanase activities; mainly in bacteria

Inverting

GH67

a-Glucuronidase and xylan a-1,2glucuronosidase activities

Inverting

GH74

Xyloglucanase and endoglucanase activities

Inverting

GH81

1,3-b-Glucanase activity

Inverting

Retaining

GH105 Rhamnogalacturonyl hydrolase

Unknown

GH113 a-Mannanase

Retaining

GH10

Endo-1,4-b-xylanase and endo-1,3-bxylanase activities in bacteria and fungi

Retaining

GH115 Xylan b-1,2-glucuronidase; b-(4-Omethyl)-glucuronidase

Inverting

GH11

Xylanase activities in bacteria and fungi

Retaining

Endoglucanase, xyloglucanase, and 1,3(4)- Retaining b-glucanase in the three domains of life

GH116 Acid a-glucosidase; a-glucosidase; axylosidase

Retaining

GH12 GH16

Broad spectrum of hemicellulases, including endo-1,3-b-glucanase, endo1,3(4)-b-glucanase, lichenanase, and xyloglucanase activities

Retaining

GH17

Glucan 1,3-b-glucosidase and lichenanase Retaining activities

GH18

Chitinases

Retaining

6

GH120 a-Xylosidase

Unknown

GH124 Endoglucanase

Inverting

GH127 a-L-Arabinofuranosidase

Retaining

GH130 1-a-D-mannopyranosyl-4-Dglucopyranose: phosphate b-Dmannosyltransferase

Inverting

223

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Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Fig. 6.4 Structures of a typical endoglucanase and exoglucanase. In each case, the structure is viewed from a perspective, which demonstrates the comparative architecture of the respective active site. Despite the sequence similarity of both enzymes and their classification as Family-6 glycoside hydrolases, their respective active-site architecture is different. The Family-6 endoglucanase (endoglucanase Cel6A from the bacterium Thermomonospora fusca, PDB code 1TML) is characterized by a deep cleft to accommodate the cellulose chain at any point along its length, whereas the active site of the Family-6 exoglucanase (cellobiohydrolase CBHI from the cellulolytic fungus, Trichoderma reesei, PDB code 1CEL) bears an extended loop that forms a tunnel, through which one of the termini of a cellulose chain can be threaded

catalysis by these enzymes progresses, it has become clear that some enzymes are capable of both endo- and exo-action (Johnson et al. 1996; Morag et al. 1991; Reverbel-Leroy et al. 1997; Sakon et al. 1997). Moreover, some glycoside hydrolase families include both endo- and exo-enzymes, again indicating that the mode of cleavage can be independent of sequence homology and structural fold. In this context, relatively minor changes in the lengths of relevant loops in the general proximity of the active site, may dictate the endo- or exo-mode of action without significant differences in the overall fold. Due to subtle but diverse chemical and structural aspects of the substrates involved, plant cell wall–degrading enzymes do not follow the same rules as common enzyme standards, such as simple proteases, DNAse, RNAse, and lysozyme. In fact, the cellulases and hemicellulases are usually very large enzymes, whose molecular masses often exceed those of proteases by factors of 2 to 5 and more. Their polypeptide chains partition into a series of functional modules and linker segments (frequently glycosylated), which together determine their overall activity characteristics and interaction with their substrates and/or with other components of the cellulolytic and hemicellulolytic system. However, the historical division of enzymes is inappropriate for the classification of the cellulases and other glycoside

hydrolases. Like other enzymes (e.g., proteases), previous classification systems of the glycoside hydrolases centered on the types of substrates and the bonds cleaved by a given enzyme. The problem with the glycoside hydrolases is that the polysaccharide substrates and particularly the bonds they cleave are all quite similar, and classification of the different types of enzymes according to conventional criteria often misses the mark. Consequently, alternative approaches were pursued. Over the past decade or so, the definitive trend has evolved to classify the different glycoside hydrolases into groups based on common sequence, structural fold and mechanistic themes (Davies and Henrissat 1995; Henrissat 1991; Henrissat and Bairoch 1996; Henrissat and Davies 1997; Henrissat et al. 1998). A comprehensive, authoritative website that provides a complete and growing catalog of the different glycoside hydrolase families is available (Coutinho and Henrissat 1999a, b, c; Coutinho et al. 2003a, b, c; Henrissat and Coutinho 2001; Henrissat et al. 2003; Cantarel et al. 2009): The CarbohydrateActive Enzymes server (http://www.cazy.org/). The website also provides similar sequence information for additional types of enzymes that participate in the degradation of plant cell wall polysaccharides, namely, carbohydrate esterases (e.g., that cleave acetyl, feruloyl and cinnamoyl groups from xylans) and polysaccharide lyases (that act on pectin). Additional associated modular components of these enzymes, particularly the carbohydrate-binding modules (CBMs), are also classified into families and documented exhaustively. An extensive list of sequenced genomes is included, which contains the carbohydrate-active enzymes encoded by the genome (‘‘CAZome’’) of the given bacterium and facilitates insight into the nature and extent of the metabolism of complex carbohydrates of the species and comparison between both related and unrelated species. The site contains excellent introductory explanatory material, and the interested reader is encouraged to use this site extensively. Moreover, a companion website, CAZypedia (http://www. cazypedia.org/index.php/Main_Page), provides an encyclopedic resource for detailed understanding of the different glycoside hydrolase families.

Cellulases The cellulases include the large number of endo- and exoglucanases which hydrolyze b-1,4-glucosidic bonds within the chains that comprise the cellulose polymer (Be´guin and Aubert 1994; Haigler and Weimer 1991; Tomme et al. 1995a). Thus, in principle, the degradation of cellulose requires the cleavage of a single type of bond. Nevertheless, in practice, we find that cellulolytic microorganisms produce a variety of complementary cellulases of different specificities from many different families. The major glycoside hydrolase families of cellulases include GH5, GH6, GH7 (found in fungi), GH8, GH9, GH12, GH44, GH45, GH48, GH74, and GH124. It may seem somewhat surprising that the combined effect of so many different enzymes is required to degrade such a chemically simplistic substrate. This complexity reflects the

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

difficulties an enzyme system encounters upon degrading such a highly crystalline substrate as cellulose. As described in the previous section, cellulases that degrade the cellulose chain can be either ‘‘endo-acting’’ or ‘‘exo-acting.’’ Moreover, the degradation of crystalline cellulose should be viewed threedimensionally and in situ, where the cellulose chains are packed within the microcrystal, thus generating the remarkably stable physical properties of the crystalline substrate. The enzymes have to bind to the cellulose surface, localize and isolate suitable chains, destined for degradation. It would seem logical that amorphous regions or defects in the crystalline portions of the substrate would be favorable sites for initiation of the process. The structural as opposed to chemical heterogeneity of the substrate dictates the synergistic action of a complex set of complementary enzymes toward its complete digestion. Various models have been suggested to account for the observed synergy between and among two or more different types of cellulases. For example, an endo-acting enzyme can produce new chain ends in the internal portion of a polysaccharide backbone, and the two newly exposed chains would then be available for action of exo-acting enzymes. In addition, two different types of exo-glucanases may exhibit different specificities by acting on a cellulose chain from opposite ends (i.e., the reducing versus the nonreducing end of the polymer). Likewise, an endoglucanase may be selective for only one of the two sterically distinct glucosidic bonds on the cellulosic surface. In addition, some cellulases may display high levels of activity at the beginning of the degradative process, i.e., on the highly crystalline material, whereas others would be selective for newly exposed, partially degraded chains, otherwise embedded within the crystal. Still others would show very high levels of activity after the degradative process has advanced, and cellulose chains which have been freed of the crystalline setting would then be hydrolyzed quite rapidly. A collection of various enzymes, which exhibit complementary specificities and modes of action, would account for the observed synergistic action of the complete cellulase ‘‘system’’ in digesting the cellulosic substrate. In addition to endo- and exo-glucanases, included in the overall group of cellulases are the b-glucosidases (EC 3.2.1.21), which hydrolyzes terminal, nonreducing b-D-glucose residues from cellooligodextrins. These enzymes are members of the following glycoside hydrolase families: GH1, GH3, and GH116. In particular, this type of enzyme cleaves cellobiose—the major end product of cellulase digestion—to generate two molecules of glucose. Some b-glucosidases are specific for cellobiose whereas others show broad specificity for other b-D-glycosides, e.g., xylobiose. Often, the b-glucosidases are associated with the microbial cell surface and hydrolyze cellobiose to glucose before, during or after the transport process. Among the novel glycoside hydrolase families, a new and important oxidative family, previously classified as a glycoside hydrolase family (GH61), was found in fungi which break internal glucan bonds (Beeson et al. 2012). A similar oxidative family of enzymes has also been proposed to exist in bacteria – the Family-33 CBMs, which were originally considered to be CBMs

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(Forsberg et al. 2011). Both the GH61 and the CBM33s will have to be reclassified separately as oxidative enzymes associated to the other CAZymes.

Hemicellulases Strictly speaking, hemicellulases are not the precise subject of this chapter, since they do not directly sever the b-1,4-glucosidic bond of cellulose. Nevertheless, in nature, they are essential to the bacterial degradation of insoluble cellulose, since the natural bacterial substrate—the plant cell wall—comprises an architecturally cogent composite of cellulose and hemicelluloses. In natural systems, the two types of polysaccharides cannot be easily separated, and microbial systems have to deal simultaneously with both. The xylan component is particularly of interest for several reasons: (a) xylan is a major hemicellulosic component of the plant cell wall; (b) the xylanases are well-defined enzymes, closely associated with the cellulase; and (c) the repeating units (both xylose and xylobiose) bear striking structural resemblance to their cellulosic counterparts (i.e., glucose and cellobiose). In contrast to cellulose degradation, the degradation of the hemicelluloses imposes a somewhat different challenge, since this group of polysaccharides includes widely different types of sugars or nonsugar constituents with different types of bonds. Thus, the complete degradation of hemicellulose requires the action of different types of enzymes. These enzymes, the hemicellulases, can differ in the chemical bond they cleave, or, as in the case of the cellulases, they may cleave a similar type of bond but with different substrate or product specificity (Biely 1985; Coughlan and Hazlewood 1993; Eriksson et al. 1990; Gilbert and Hazlewood 1993). Hemicellulases can be divided into two main types, those that cleave the main chain backbone, i.e., xylanases or mannanases, and those that degrade side chain substituents or short end-products, such as arabinofuranosidase, glucuronidase, acetyl esterases, and xylosidase. Like the cellulases, hemicellulases can be of the endo or exo types. A schematic view of the types of bonds that would be hydrolyzed by different types of hemicellulases is presented in > Fig. 6.3.

Xylan-Degrading Enzymes The xylanases are by far the most characterized and studied of the hemicellulases and involve the cleavage of a major main chain backbone. Endoxylanases (1,4-b-D-xylan xylanhydrolase, EC 3.2.1.8) hydrolyze the 1,4-b-D-xylopyranosyl linkage of xylans, such as D-glucurono-D-xylans and L-arabinoD-xylan. These single-subunit enzymes from both fungi and bacteria exhibit a broad range of physiochemical properties, whereby two main classes have been described: alkaline proteins of low Mr ( Table 6.3). Like the other CAZymes, the PLs are modular proteins, which serve to complement the activities of the cellulases and other enzymes to better degrade the plant cell wall polysaccharide components.

Carbohydrate Esterases The side chain substituents of xylan are composed not only of sugars but also of acidic residues, such as acetic, ferulic (4-hydroxy-3-methoxycinnamic), or p-coumaric (4-hydroxycinnamic) acids. Carbohydrate esterases that cleave

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Table 6.3 Polysaccharide lyase families involved in the degradation of plant cell wall polysaccharides and their enzymatic activities (See CAZy website for more details: http://www.cazy.org/) PL family

Enzymes

. Table 6.4 Carbohydrate esterase families involved in the degradation of plant cell wall polysaccharides and their enzymatic activities (See CAZy website for more details: http://www.cazy.org/) CE family

Enzymes

PL1

Pectate lyase, exo-pectate lyase and pectin lyase activities

CE1

Acetyl xylan esterase, cinnamoyl esterase, feruloyl esterase and carboxylesterase activities

PL3

Pectate lyases

CE2

Acetyl xylan esterases

PL9

Pectate lyase and exopolygalacturonate lyase activities

CE3

Acetyl xylan esterases

PL10

Pectate lyases

CE4

Acetyl xylan esterases

PL11

Rhamnogalacturonan lyase activities

CE6

Acetyl xylan esterases

CE7

Acetyl xylan esterases

CE8

Pectin methyl esterases

CE12

Pectin acetyl esterase, rhamnogalacturonan acetyl esterase, and acetyl xylan esterase activities

CE15

4-O-Methyl-glucuronoyl methyl esterase activity

> Fig.

these residues (see 6.3) are found in enzyme preparations from both hemicellulolytic and cellulolytic cultures (Borneman et al. 1993). Such enzymes sometimes represent separate modules, separated by linker segments from other cellulolytic or hemicellulolytic catalytic modules in the same polypeptide chain. Like the glycoside hydrolases, the carbohydrate esterases are classified into families according to sequence homology and common structural fold (http://www.cazy.org/) and they frequently appear together with other modular components, notably xylanases from glycoside hydrolase families 10 and 11, on the same polypeptide chain. A list of the important CE families is given in > Table 6.4. Most of the families contain enzymes that exhibit acetyl xylan esterase activity. Family CE1 also has members that cleave cinnamoyl and feruloyl bonds. These enzymes are very important ones, since this would allow a bacterium to sever the xylan components that are attached covalently to lignin. As lignin and its degradation products are frequently deleterious to enzymes that degrade plant-derived polysaccharides and to the bacterium itself, the action of the ferulic and coumaric acid esterases would promote more effective degradation of the xylan upon its separation from the lignin.

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Teeri et al. 1992). Each enzyme contains at least one catalytic module, which catalyzes the actual hydrolysis of the glycosidic bond and provides the basis for classification of the simple enzymes (i.e., those containing a single catalytic module). Other accessory or ‘‘helper’’ modules assist or modify the primary hydrolytic action of the enzyme, thus modulating the overall properties of the enzyme. Some of the different themes illustrating the modular compositions of the cellulases and related enzymes are presented in > Fig. 6.5. In many cases, certain patterns can be observed between the catalytic module(s) and the types of ancillary modules, notably the CBMs, which consistently occur in natural enzyme systems of the cellulolytic bacteria. Knowledge of the different modular components that comprise a given enzyme and thus modulate its activity can thus suggest the functional characteristics of the enzyme.

The Catalytic Modules: Families of Enzymes

Cellulases and Hemicellulases Are Modular Enzymes The initial contribution of biochemical methods for determining the characteristics of a given cellulase was extended immeasurably by the contribution of molecular biology and bioinformatics. By comparing the sequences of the cellulases and related enzymes, an entirely new view of these enzymes emerged. Cellulases and hemicellulases are composed of a series of separate modular components. This fact explains the very large size of some of these enzymes and gives us some insight into their complex mode of action. Each module comprises a consecutive portion of the polypeptide chain and forms an independently folding, structurally and functionally distinct unit (Coutinho and Henrissat 1999a, b, c; Gilkes et al. 1991;

The definitive component of a given enzyme is the catalytic module. Former EC-based classification schemes according to substrate specificity are now considered somewhat obsolete, since they fail to take into account the structural features of the enzymes themselves and for the compound reasons listed in the previous sections. The catalytic modules of glycoside hydrolases are now categorized into families according to amino acid sequence homology (Cantarel et al. 2009; Coutinho and Henrissat 1999a, b, c; Henrissat 1991; Henrissat and Bairoch 1996; Henrissat and Davies 1997; Henrissat et al. 1998). For more information, see the website server for the CarbohydrateActive Enzymes (CAZy), designed and maintained by Bernard Henrissat and Pedro Coutinho (http://www.cazy.org/). The enzymes of a given glycoside hydrolase family are similar in sequence, they display the same structural topology, and the positions of the catalytic residues are conserved with respect to the

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an overall retention or an inversion of the configuration of the anomeric carbon (Davies and Henrissat 1995; McCarter and Withers 1994; White and Rose 1997; Withers 2001). In both cases, cleavage is catalyzed primarily by two active-site carboxyl groups. One of these acts as a proton donor and the other as a nucleophile or base. Retaining enzymes function via a doubledisplacement mechanism, by which a transient covalent enzyme-substrate intermediate is formed (> Fig. 6.6a). In contrast, inverting enzymes employ a single-step mechanism as shown schematically in > Fig. 6.6b. The distance between the acid catalyst and the base represents the major structural difference between the two mechanisms. In retaining enzymes, the distance between the two catalytic residues is about 5.5 A˚, whereas in inverting enzymes, the distance is about 10 A˚. In the inverting enzymes, additional space is provided for a water molecule, involved directly in the hydrolysis, and the resultant product exhibits a stereochemistry opposite to that of the substrate. In all cases, the mechanism of hydrolysis is conserved within a given glycoside hydrolase family (Coutinho and Henrissat 1999a, b, c; Davies and Henrissat 1995; Henrissat and Davies 1997). . Fig. 6.5 Scheme illustrating the diversity of the modular architecture of cellulases and other glycoside hydrolases. The different modules are grouped into families according to conserved sequences as shown by the pictograms in the Figure. (a) One of the most common types of cellulases consists of a catalytic module, flanked by a CBM at its N- or C-terminus. The particular enzyme shown in (a) comprises a catalytic module from Family-48 and a Family-2 CBM. (b) Cellulosomal enzymes are characterized by a ‘‘dockerin module’’ attached to a catalytic module. In this case, the same type of enzyme as in (a), carrying a Family-48 catalytic module, harbors a dockerin module instead of a CBM. (c) Many cellulases contain ‘‘X domains,’’ i.e., domains of unknown (as yet undefined) function. Often such domains prove to be a CBM when the appropriate binding specificity is determined experimentally. (d) Some enzymes have more than one CBM. Often, one CBM, such as the Family-3 CBM shown in the Figure, serves to bind the cellulase strongly to the flat surface of the insoluble substrate, whereas the other one (the Family-3c CBM) acts in concert with the catalytic module by binding transiently to a single cellulose or to a hemicellulose chain. (e) Some cellulosomal cellulases have a CBM together with a dockerin in the same polypeptide chain. (f) Some cellulases have more than one type of catalytic module, such as the Family-5 and Family-44 modules shown in the Figure, and the two probably work in concerted fashion to degrade the substrate efficiently

common fold. X-ray crystallography has provided a general overview of the structural themes of the glycoside hydrolases and their interaction with their intriguing set of substrates (Bayer et al. 1998; Davies and Henrissat 1995; Henrissat and Davies 1997). The mechanism of cellulose and hemicellulose hydrolysis occurs via general acid catalysis and is accompanied by either

Carbohydrate-Binding Modules (CBMs) In addition to the catalytic module, free cellulases and hemicellulases usually contain at least one carbohydrate-binding module (CBM) as an integral part of the polypeptide chain (Linder and Teeri 1997; Tomme et al. 1995b). The CBM serves predominantly as a targeting agent to direct and attach the catalytic module to the insoluble crystalline substrate. Like the catalytic modules, the CBMs are categorized into a series of families according to sequence homology and consequent structural fold. For historical reference, until the year 2000 or so, the original term used in the literature for such substrate-binding modules was CBD, as an indication of cellulose-binding domain. However, CBD is deceptive, since not all of them bind to cellulose, and some families have members that bind to cellulose as well as other types of polysaccharides. It became clear that a more general term was required, and the term CBM (carbohydrate instead of cellulose, module instead of domain) was chosen and is clearly more appropriate on both counts. Some CBM families (or subfamilies or family members) bind either preferentially or additionally to other insoluble polysaccharides, e.g., xylan or chitin. For example, the Family-5 CBM and some of the members of the Family-3 CBMs bind to chitin as well as cellulose (Brun et al. 1997; Morag et al. 1995). Moreover, the Family-2 CBMs can be divided into two subfamilies, one of which indeed binds preferentially to insoluble cellulose, but the other binds to xylan (Boraston et al. 1999). The molecular basis for this was proposed to reflect the fact that in the first subfamily, three surface-exposed tryptophans contribute to cellulose binding (Simpson et al. 1999; Williamson et al. 1999). However, in the case of the xylan-binding members, one of these tryptophans is missing, whereas the other two

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

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. Fig. 6.6 The two major catalytic mechanisms of glycosidic bond hydrolysis. (a) The retaining mechanism involves initial protonation of the glycosidic oxygen via the acid/base catalyst with concomitant formation of a glycosyl-enzyme intermediate through the nucleophile. Hydrolysis of the intermediate is then accomplished via attack by a water molecule, resulting in a product which exhibits the same stereochemistry as that of the substrate. (b) The inverting mechanism involves the single-step protonation of the glycosidic oxygen via the acid/base catalyst and concomitant attack of a water molecule, activated by the nucleophile. The resultant product exhibits a stereochemistry opposite to that of the substrate. The type of mechanism is conserved within a given glycoside hydrolase family and dictated by the active-site architecture and atomic distance between the acid/base and nucleophilic residues (aspartic and/or glutamic acids)

assume a different conformation, thereby allowing them to stack against the hydrophobic surfaces of two xylose rings of a xylan substrate. Other types of CBM prefer less crystalline substrates (e.g., acid-swollen cellulose), single cellulose chains, and/or soluble oligosaccharides, e.g., laminarin (1,3-b-glucan) and barley 1,3/1,4-b-glucan (Tomme et al. 1996a, b; Zverlov et al. 2001).

Still others exhibit alternative accessory function(s), a topic to be described below in more detail. Moreover, the CBMs responsible for the primary binding event may further disrupt hydrogen-bonding interactions between adjacent cellulose chains of the microfibril (Din et al. 1994), thereby increasing their accessibility to subsequent attack by the hydrolytic module.

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Consequently, the concept of CBM has been broadened and redefined as CBM—i.e., carbohydrate-binding module (Boraston et al. 1999, 2004; Coutinho and Henrissat 1999a, b, c). In the previous edition of The Prokaryotes, more than a decade ago, 26 different CBM families were described. To date (March 2012), the number of CBM families have increased to 64 (http://www.cazy.org/). A CBM can be identified as a member of the family on the basis of sequence and position of binding residues before the binding function itself is established; nevertheless, it is imperative to confirm experimentally the specificity of binding of an individual CBM. The structures of CBMs from a number of families and subfamilies have been determined, and an understanding of their structures has provided interesting information regarding the mode of binding to cellulose. Those that bind to crystalline substrates, appear to do so via a similar type of mechanism. One of the surfaces of such CBMs is characteristically flat and appears to complement the flat surface of crystalline cellulose. A series of aromatic amino acid residues on this flat surface form a planar strip (Mattinen et al. 1997; Simpson and Barras 1999; Tormo et al. 1996; Koyama et al. 1997; Lehtio et al. 2003; Ding et al. 2006) that stack opposite the glucose rings of a single cellulose chain. In addition, to the planar aromatic strip, several polar amino acid residues on the same surface appear to anchor the CBM to two adjacent cellulose chains. The binding of the CBM to crystalline cellulose would thus involve precisely oriented, contrasting hydrophobic and hydrophilic interactions between the reciprocally flat surfaces of the protein and the carbohydrate substrate. Together they provide a selective biological interaction, which contributes to the specificity that a CBM exhibits toward its structure. In some cases, the putative binding surface turns out to be irregular instead of flat, which may obstruct binding (Petkun et al. 2010). In contrast to the interaction with the crystalline cellulose surface, other CBMs seem to interact with single cellulose chains. The Family-3c and Family-4 CBMs preferentially bind to non-crystalline forms of cellulose and clearly have a different function in nature (Johnson et al. 1996; Sakon et al. 1997; Tomme et al. 1996a, b). For example, the role of Family-4 CBM may be to recognize, bind to, and deliver an appropriate catalytic module to a cellulose chain, which has been loosened or liberated from a more ordered arrangement within the cellulose microfibril. The binding of the Family-3c CBM to single cellulose chains and its remarkable role in cellulose hydrolysis will be discussed later (> Fig. 6.9: section on > ‘‘Helper Modules’’). The role of the CBMs has been expanded recently from the conventional substrate targeting function to cell-surface attachment (Ezer et al. 2008; Montanier et al. 2009) and vital important biomass sensing functions leading to transcriptional regulation (Kahel-Raifer et al. 2010; Nataf et al. 2010; Bahari et al. 2011).

The Family-9 Cellulases: An Example This section pertains to enzyme diversity and how a single type of catalytic module can be modified by the class of helper

module(s) that flank its C- or N-terminus. We are only at the beginning in our understanding of how the modular arrangement affects the overall activity and function of a given enzyme. In its simplest form, an enzyme would presumably consist of a single catalytic module, usually with a standard CBM, which would target the enzyme to the crystalline substrate. Indeed, this is the norm for many individual glycoside hydrolase families. However, in others, e.g., the Family-9 cellulases, the catalytic modules commonly occur in tandem with a number of accessory modules. Although the story remains rather incomplete, we can discuss the currently available information regarding Family 9 and draw several interesting conclusions from the few publications on this currently developing subject. Family-9 Theme and Variations

The crystal structure of the Family-9 catalytic module displays an (a/a)6-barrel fold and inverting catalytic machinery. There are numerous Family-9 cellulases of plant origin (Coutinho et al. 2003a, b, c) the great majority of which are lone catalytic modules that lack accessory modules. Another type of eukaryotic Family-9 cellulase that lacks helper modules is produced by the termite (Ni et al. 2005). Only a few of the prokaryotic Family-9 enzymes consist of a solitary catalytic module (> Fig. 6.7a). The prokaryotic Family-9 enzymes, however, are almost invariably decorated with a variety of subsidiary modules that modulate the activity of the catalytic module. Microbial Family-9 cellulases commonly conform to one of the themes shown in > Fig. 6.7, which were recognized in the previous edition. In one of these, the catalytic module is followed immediately downstream by a fused Family-3c CBM (> Fig. 6.7b). This particular type of CBM imparts special characteristics to the enzyme (see below). A second theme consists of an immunoglobulin-like (Ig) domain (of unknown function) immediately upstream to the catalytic module (> Fig. 6.7c). A variation of the latter theme includes a Family-4 CBM at the N-terminus of the enzyme, followed by an Ig domain and Family-9 catalytic module (> Fig. 6.7d). In addition to the above-described modular arrangement, each of the free prokaryotic enzyme systems includes a standard CBM that binds strongly to crystalline cellulose. In the last decade, several additional themes have been described, notably GH9-CBM3c0 CBM3b0 (i.e., a GH9 catalytic module followed by two successive subtypes of CBM3) with a C-terminal dockerin. This theme is present in the genomes of Clostridium thermocellum, Acidothermus cellulolytics, and Clostridium clariflavum. The Family-9 glycoside hydrolase of the cellulosomal scaffoldin from the cellulolytic anaerobic bacterium Acetivibrio cellulolyticus contains no helper module (Ding et al. 1999). The A. cellulolyticus enzyme forms part of a multi-modular scaffoldin, but the catalytic module appears to be a functionally distinct entity that lacks adjoining helper modules. The other modules are conventional scaffoldin-associated modules, e.g., cohesins and a true cellulose-binding CBM. This thematic arrangement of the Family-9 cellulases is mirrored in the respective sequences of the catalytic modules. The divergent sequences are reflected by the phylogenetic

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

6

. Fig. 6.7 Theme and variations: schematic view of some of the modular arrangements of the Family-9 glycoside hydrolases. (a) The solitary catalytic module; (b) the catalytic module and fused Family-3c CBM; (c) immunoglobulin-like (Ig) domain, fused to the catalytic module; (d) successive Family-4 CBM, Ig, and catalytic modules. The representations of the different modules are based on their known structures and are presented sequentially, left-to-right, from the N- to C-terminus. Structures in (a) and (b) are derived from cellulase E4 from Thermomonospora fusca (PDB code, 1TF4), those in (c) and (d) are from the CelD endoglucanase of C. thermocellum (PDB code, 1CLC). The Figure used for the Family-4 CBM in (d) is derived from the nmr structure of the N-terminal CBM of Cellulomonas fimi b-1,4-glucanase CenC (PDB code, 1ULO). The structures in (b) and (c) are authentic views of the respective crystallized bi-modular protein components. The CBM in (d) has been placed manually to indicate its N-terminal position in the protein sequence, but its spatial position in the quaternary structure and the structure of the linker segment remains unknown

relationship of the parent cellulases (> Fig. 6.8). Thus, the simplest cellulases (the Group A eukaryotic cellulases from plants) that lack adjacent helper modules are all phylogenetically related (Theme A). Interestingly, the catalytic module of ScaA from A. cellulolyticus is distinct from the other groups designated in > Fig. 6.8, but closest to the plant enzymes, as might be anticipated from its lack of a helper module. In a similar manner, catalytic modules from cellulases that are fused to a Family-3c CBM (Group B), all map within the same branch (Theme B). On the other hand, the catalytic modules that bear an adjacent Ig-like domain all fall into a cluster on the opposite side of the tree. Cellulases which have the Ig-like domain only (Theme C) occupy a small separate branch and those that also include a Family-4 CBM (Theme D) that develops distally to form a separate subcluster.

architecturally distinct—the T. fusca Cel9A cellulase being an example of a Theme B Family-9 enzyme (see > Figs. 6.7b and > 6.8) and the C. thermocellum Cel9D cellulase being a Theme C enzyme. Fortunately, in both cases, one of the neighboring modules co-crystallized with the catalytic module, thus providing primary insight into their combined structures. In the case of T. fusca Cel9A, the catalytic module and neighboring Family-3c CBM were found to be interconnected by a long, rigid linker sequence, which envelops about half of the catalytic module until it connects to the adjacent CBM (> Fig. 6.9a). In contrast, in the C. thermocellum Cel9D, the catalytic module is adjoined at its N-terminus by a 7-stranded immunoglobulin-like (Ig) domain of unknown function. The comparison between the E4 and CelD cellulases indicates that a given type of catalytic module can be structurally and functionally modulated by different types of accessory modules.

Family-9 Crystal Structures

Crystal structures of Family-9 cellulases have been elucidated, representing two subtypes of this particular family of glycoside hydrolase. These are cellulase E4 (or Cel9A) from Thermobifida fusca (Sakon et al. 1997) and Cel9D from Clostridium thermocellum (Juy et al. 1992). These two examples are

Helper Modules

The Family-3c CBM is special. To date, this particular type of CBM has been found in nature associated exclusively with the Family-9 catalytic module. Structurally, the CBM is homologous to the other Family-3 CBMs, but contains substitutions in many

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. Fig. 6.8 Phylogenetic analysis of the N-terminal Family-9 catalytic module of ScaA and its relationship with other Family-9 members. The various theme groupings roughly follow the groups shown in > Fig. 6.7. Theme A (Group A) enzymes lack associated helper modules. Theme B (Group B) enzymes carry a fused Family-3c CBM downstream to the catalytic module. Theme C (Group C) and Theme D (Group D) enzymes carry an Ig domain upstream to the catalytic module, the Theme D enzymes having an additional N-terminal Family-4 CBM. Theme A enzymes: ScaA Acece, ScaA scaffoldin from the cellulolytic bacterium A. cellulolyticus (AF155197); and plant (eukaryotic) cellulases from Prunus persica (X96853), Populus alba (D32166), Citrus sinensis (AF000135), Persea americana (M17634), Pinus radiata (X96853), Arabidopsis thaliana (X98543), Phaseolus vulgaris (M57400), Capsium annuum (X97189), Lycopersicon esculentum (U20590). Theme B enzymes: CelF Clotm, endoglucanase F from Clostridium thermocellum (X60545); CelZ Closr, exoglucanse Z from Clostridium stercorarium (X55299); CelA Calsa, cellulase A from Caldocellum saccharolyticum (L32742); CelG Cloce, endoglucanase G from Clostridium cellulolyticum (M87018); CelI Clotm, endoglucanase I from Clostridium thermocellum (L04735); CelB Celfi, endoglucanase B from Cellulomonas fimi (M64644); E4 Thefu, endo/exoglucanase E4 from Thermomonospora fusca (M73322). Theme C enzymes: CelJ Clotm, cellulase J from Clostridium thermocellum (D83704); CelD Clotm, endoglucanase D from Clostridium thermocellum (X04584); CelC Butfi, endoglucanase C from Butyrivibrio fibrisolvens (X55732). Theme D enzymes: CbhA Clotm, cellobiohydrolase A from Clostridium thermocellum (X80993); CelA Psefl, endoglucanase A from Pseudomonas fluorescens (X12570); CelC Celfi, endoglucanase C from Cellulomonas fimi (X57858); CelI Strre, endoglucanase I from Streptomyces reticuli (X65616); E1 Thefu, endoglucanase E1 from Thermomonospora fusca (L20094). The analysis of the designated catalytic modules was performed using GenBee, based on the respective GenBank sequences (accession codes in parentheses)

important surface residues. The three-dimensional crystal structure of the T. fusca Cel9A cellulase revealed the close interrelationship between the Family-9 catalytic module and the Family3c CBM, thus suggesting a functional role as a helper module. This CBM seems not to bind directly to crystalline cellulose but appears to act in concert with the catalytic module by binding transiently to the incoming cellulose chain, which is then fed into the active-site cleft pending hydrolysis (> Fig. 6.9b) (Gal et al. 1997a; Irwin et al. 1998; Sakon et al. 1997).

The information derived from the Family-9 enzymes suggests that the activity of catalytic modules can be modulated by accessory modules. The accessory modules can either supplement or otherwise alter the overall properties of an enzyme (Bayer et al. 1998b, c). The recurrent appearance in nature of a given type of module adjacent to a specific type of neighboring catalytic module may indicate a functionally significant theme. These observations raise the possibility of a more selective role for certain types of CBM and other modules, whereby their

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complexation, the GH9 essentially lacked activity. Physical association of the two modules was shown to recover 60–70 % of the intact Cel9I endoglucanase activity.

Cellulase Analysis

. Fig. 6.9 Structural aspects of Family-9 Theme-B cellulase E4 from T. fusca. (a) ‘‘Side view’’ of the E4 molecule. Shown are the Family-9 catalytic module (turquoise, on the left), the Family-3c CBM (in yellow, on the right) and the intermodular linker (dark blue strip). The presumed path of a single cellulose chain, from the CBM to the catalytic module, is shown at the bottom of the structure (arrows). The enzyme also possesses a fibronectin-like domain (FN3) and a cellulose-binding Family 2 CBM (not shown). Note that the linker appears to serve a defined structural role by which the Family-3c CBM is clamped tightly to the catalytic module. Selected surface residues on the catalytic module along the interface of both the linker and the CBM3c also serve to fasten both features tightly to the catalytic module. (b) ‘‘Bottom view’’ of the E4 molecule (90 rotation of a). From this perspective, the proposed catalytic residues (red), positioned in the active site cleft, are clearly visible. The path of the cellulose chain (arrows) passes through a succession of polar residues (green) on the bottom surface of the CBM which would conceivably bind to the incoming cellulose chain and serve to direct it toward the active-site acidic residues of the catalytic module

association with certain types of catalytic modules could signify a ‘‘helper’’ role. The helper module would provide hydrolytic efficiency and alter the catalytic character of the enzyme. Interestingly, in recent work on a Theme B enzyme, Cel9I from C. theromcellum (Burstein et al. 2009), recombinant forms of the individual GH9 catalytic module and CBM3c (together with the intermodular linker) were expressed individually, and the two modules underwent self-assembly to form a complex. Before

The biochemical characterization of cellulases is in many cases a difficult task owing to the large variety of enzyme types and modes of action. At first glance, it is an intriguing phenomenon that for such a simple reaction (i.e., the hydrolysis of the b-1,4glucose linkage in a linear glucose chain), nature has evolved so many types of cellulases. The vast varieties of enzymes are found not only among the different species of cellulolytic bacteria but also within the same organism. The reason for this extensive diversity comes from the insoluble nature of cellulose and the fact that although the chemical composition of the homopolymer is rather trivial, the physical and three-dimensional arrangement of the chains within the crystalline and amorphous regions of the microfibril can differ significantly. Regarding the enzymes that degrade the substrate, the modular nature of the cellulases contributes additional degrees of complexity in our quest to characterize a given enzyme. Thus, the number, types, and arrangement of the accessory modules vis-a`-vis the catalytic module are important structural features that modulate the overall activity of the enzyme in question. This descriptive information should always be defined for a recombinant enzyme. Whenever possible, it is desirable to determine the relative contribution of the individual accessory modules to the activity of the enzyme. In this regard, the affiliation of a given module, e.g., CBM, into a defined family does not necessarily define its contribution to enzyme activity, as different specificities and functions have been attributed to different members of the same family of module. Moreover, sequences for the different ‘‘X’’ modules (i.e., modules for which the function remains undefined) are widespread, most of which probably play a carbohydrate binding or processing role in assisting the catalytic module(s) in its capacity to hydrolyze the substrate. Two decades ago, the range of cellulases and hemicellulases within a given species was assessed mainly by biochemical techniques. In some cases, individual enzymes were isolated and their properties assessed using desired insoluble or soluble substrates. Another approach involved electrophoretic separation of cell-derived or cell-free extracts, and analysis of desired activities using zymograms. There are advantages and disadvantages with each of these strategies, and the employment of combined complementary approaches is always advisable. Molecular biology techniques are also used to reveal cellulase and hemicellulase genes, which can often be characterized on the basis of sequence homology with related, known genes (Be´guin 1990; Hazlewood and Gilbert 1993) or according to their GH family membership (> Table 6.2). If further information is required on the structure or action of a given enzyme, the gene can then be expressed in an appropriate host organism, and the properties of the product can be characterized.

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It is always instructive to compare the properties of an expressed gene product with those of the same protein isolated from the original bacterial culture. The results may be surprising; there are hazards inherent to both approaches. Expression of a gene may yield preparations with reduced or altered enzymatic properties. In this context, the expressed gene product may not have been folded properly. It is of course assumed that the investigator has taken the time and trouble to sequence the cloned gene to ensure no mutations have occurred. Unlike a gene expressed in a host cell environment, the native counterpart may have undergone posttranslational modifications (e.g., glycosylation, proteolytic truncation, etc.) that improve its physicochemical properties. Moreover, since the cellulase system in the native environment includes numerous enzyme types, often exhibiting similar molecular masses and other physical characteristics, the reputed purification of a given extracellular cellulase may still include contaminating enzymes that alter (usually increasing greatly due to synergistic action of two or more enzymes) the true enzymatic properties of the desired enzyme. The onus belongs to the conscience of the investigating scientist when publishing the properties of a given enzyme. Too often, erroneous data that enter the scientific literature are taken as fact. One should particularly be wary of comparing enzymatic activities of the same or similar types of enzymes (e.g., members of the same family) that have been published at different times and by different laboratories. During the past decade, the phenomenal decrease in the costs associated with genomic and metagenomic sequencing efforts has completely altered accepted methodology for enzyme discovery. Today, sequencing of a cellulolytic microbe with concomitant bioinformatic annotation yields dozens and sometimes hundreds of new enzymes which can generally be included into the known families of glycoside hydrolases. The gargantuan efforts in establishing the CAZy database (http://www.cazy.org/) (Cantarel et al. 2009) have today provided the informed researcher with tools to determine the general features of a given enzyme. Nevertheless, researchers who seriously seek to understand more deeply the action of a newly discovered enzyme must perform the biochemical, structural, and enzymological studies in a meticulous manner. The establishment of novel families – i.e., glycoside hydrolases as well as other carbohydrate-active enzyme superfamilies, requires much more intensive and elegant studies of this nature. This is particularly evident for many types of cellulases, where no simple colorimetric assays exist. In some cases, chromogenic substrates or assays are available and the detection of cellulolytic assays are, in this case, more straightforward; in others, the activity is much more subtle. This is reflected in the fact that most of the known glycoside hydrolase families which include genuine cellulase were identified early on, since their members were identified colorimetrically. Since that time, new families of cellulases were difficult to establish, mainly due to the lack of a simple comprehensive assay or sets of assays that would definitively identify new types of cellulolytic activity. In the early 1990s, an important family was

discovered that includes exoglucanases (glycoside hydrolase Family 48). The founding member of this family was a predominant component of the C. thermocellum cellulosome (Morag et al. 1991; Wang et al. 1993, 1994). Subsequent research has established that a member of this family is consistently a major component of each newly discovered cellulosome. In addition, members of this family have been discovered in both free and multifunctional cellulases. Nearly two decades then passed until a new type of cellulase was discovered (Bra´s et al. 2011), which allowed formation of a new family (glycoside hydrolase Family 124). In this case, the actual cellulose-degrading function was somewhat cryptic and its detection required a combined approach until the enzyme could be verified as a cellulase. Clearly, with continuing genomic and metagenomic sequencing, there are myriads of unknown and novel types of cellulases and other associated plant-derived polysaccharidedegrading enzymes that await future discovery. Novel, preferably medium- or high-throughput approaches will be required to promote this endeavor. The assessment of cellulase activity is indeed a complicated undertaking, and there is no clear or standard methodology for doing so. This predicament apparently reflects a combination of factors, including the complex nature of the substrate, the multiplicity of enzymes and their synergistic action, and the variety of products formed. The fact that cellulose is an insoluble substrate converted to lower-order cellooligosaccharide products is a further complication. It must be noted that as the cellooligomers increase in length, they become less soluble, such that cellooctaose of eight glucose units is no longer soluble in aqueous solutions. Moreover, the accumulation of one (particularly cellobiose) or more of the cellulose degradation products may be inhibitory toward enzymatic activity. Today, the study of cellulase action usually includes, in addition to conventional biochemical assays, the analysis of the primary structure and the assignment of the various modules into known families. The catalytic modules can usually be assigned into one of the known glycoside hydrolase families (Henrissat and Bairoch 1996; Henrissat and Davies 1997). Whenever the sequence of a known polysaccharide-degrading enzyme failed to match a known family, a new family of glycoside hydrolase was established. This approach was extensively developed in the last decade, due to the increasing number of available DNA sequences and bioinformatics analysis tools. At the same time, an increasing number of crystal or solution structures of various catalytic and accessory modules were published that allow us to examine a new protein sequence in light of its structure. Sometimes, the publication of the structure of an accessory module precedes determination of its function. We can divide the analysis of a newly described prospective cellulase into several stages, such that a variety of complementary approaches are currently in use in order to classify the enzyme. Some of the questions one may ask are: 1. What is the primary structure (the amino acid sequence) of the enzyme? What are the binding residues and/or binding

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

2.

3. 4. 5.

module(s) associated with the enzyme? What are its other accessory modules and their respective role(s) in catalysis or stability? Is the enzyme a ‘‘true’’ cellulase, i.e., is its preferred substrate a cellulose or a cellulose-degradation product, and can the enzyme act alone on insoluble cellulose. This is to be distinguished from simple endoglucanases and exoglucanases and their activities on model substrates. What is the mode of action? Does the enzyme act as an endoglucanase, an exoglucanase, or a processive enzyme? What is the stereochemistry of the reaction? Does the enzyme exhibit an inverting or retaining mechanism? What are the catalytic residues: the acid/base residue and the nucleophile that characterize a glycoside hydrolase?

In the early years of cellulase research, several extensive reviews and book chapters dealt with different assays of cellulose degradation (Ghose 1987; Wood and Kellogg 1988). In this treatise, we will briefly summarize the various approaches currently in use and direct the reader to the relevant literature. While characterizing the activity of a new enzyme preparation, one has to bear in mind several secondary or indirect issues, such as the purity of the protein preparation, the sensitivity of the assay used, and the cross-reactivity of the expected enzymatic activities. In some cases, only detailed kinetic analysis can provide appropriate characterization of the enzyme. As for many other types of glycoside hydrolases, cellulases can exhibit cross-reactivity with substrates of similar structure. This is particularly true when using, e.g., p-nitrophenyl derivatized substrates that provide highly sensitive assays. However, in many cases, such a soluble synthetic chromogenic substrate can fit the active-site pocket of a related but atypical enzyme, which catalyzes its hydrolysis. For example, Family-10 glycoside hydrolases are typically xylanases but individual members of this family can readily hydrolyze p-nitrophenyl cellobioside which is a typical cellulase substrate. Without a detailed comparative kinetic analysis (kcat/km) using different substrates, the true specificity of the enzyme might be overlooked. Today, given the amino acid sequence of the protein, its assignment to a given glycoside hydrolase family can in many cases provide a reasonable general indication of its activity. The description of the modular structure provides additional knowledge that can imply how the catalytic function might be modulated, but this knowledge can also be misleading. In the final analysis, there is no substitute for extensive biochemical and biophysical characterization of the given protein (recombinant or native) and its catalytic properties. In case of a native enzyme, it is imperative to ensure that contaminating enzymatic activities have been removed. This is not a trivial undertaking. In the case of a recombinant form of an enzyme, it is imperative to ensure that the enzyme is correctly folded and the activity(ies) is (are) indicative of the parent protein. In the case of multi-modular enzymes, wherein the ancillary modules may alter the character of the catalytic module, again, these efforts are nontrivial. General procedures for assaying for cellulase and hemicellulase activities are very well documented in the

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Methods in Enzymology Volume 160 (Wood and Kellogg 1988) and a new Volume of this series is forthcoming (Gilbert, 2012). Conventional procedures for cellulase assay have been defined precisely by IUPAC (Ghose 1987). However, due to the complexity of the substrate and enzyme systems, these procedures can only provide a starting point for understanding the true nature of the enzyme in question. Since the publication of Part A of this treatise (Coughlan and Mayer 1992), many of the previously reported assays of cellulase activity are still in common use. These include the use of soluble, derivatized forms of cellulose, e.g., carboxymethyl cellulose and hydroxymethyl cellulose, as conventional substrates for determining endoglucanase activity. In addition, a derivatized, colored form of insoluble cellulose, i.e., azure cellulose, is frequently used as an indication of cellulase activity. Zymograms with such colored embedded substrates are useful in detecting endoglucanase or xylanase activities (Be´guin 1983). Individual soluble cellooligomers (cellotetraose, cellopentaose, cellohexaose, etc.) are still used as substrates for analyzing enzyme action, but the reliance on these substrates as determinants for assessing cellulase activity is no longer a definitive approach. Substrate analogues and reagents were developed that include the use of thioglycoside substrates (Driguez 1997), fluoride-derivatized sugars (Williams and Withers 2000), and chromophoric and fluorescent cellooligosaccharides (Claeyssens and Henrissat 1992; O’Neill et al. 1989; van Tilbeurgh et al. 1985). An ultraviolet-spectrophotometric method and an enzyme-based biosensor have also been described (Bach and Schollmeyer 1992; Hilden et al. 2001). In addition, a novel and intriguing bifunctionalized fluorogenic tetrasaccharide has been developed as an effective reagent for measuring the kinetic constants of cellulases by resonance energy transfer (Armand et al. 1997). The thiooligosaccharides serve as competitive inhibitors that mimic natural substrates but are enzyme resistant (Driguez 1997). In this type of oligosaccaride, the oxygen of a bond to be cleaved is replaced by sulfur. The thiooligodextrins are sometimes more soluble than the native cellodextrins and longer chains can be synthesized. The modified sugars can be used in biochemical studies or crystallographic studies to gain some information about the geometry of the active site or determine the mechanism of action of an enzyme.

Determination of ‘‘True’’ Cellulase Activity: Solubilization of Crystalline Cellulose Substrates True cellulase activity is usually defined as the ability to solubilize to an appreciable degree insoluble, ‘‘crystalline’’ forms of cellulose. The extent of hydrolysis can be evaluated by turbidity assays, weight loss of insoluble material, generation of reducing power, and accumulation of soluble sugars. It is important to realize that crystalline cellulose is not of uniform composition and therefore the rate of catalysis is in most cases not linear with time or enzyme concentration. Notably, the different

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preparations of crystalline cellulose contain varying levels of loosely associated loops and chains. The latter are readily accessible to hydrolysis by a given enzyme and lead to relatively high initial rates of activity, which do not reflect the actual degree of true cellulase activity. For example, such loose chains can be degraded by a relatively ineffectual enzyme, whereas the crystalline portions of the substrate will be immune to further hydrolysis by the same enzyme. To overcome these difficulties, IUPAC suggests determining the amount of enzyme required to achieve digestion of 5.2 % of the insoluble substrate (e.g., filter paper) in 16 h (Ghose 1987; Irwin et al. 1993). Cellulose substrates commonly in use include Avicel, filter paper, cotton, Solka Floc, as well as bacterial cellulose from Acetobacter aceti and algal cellulose prepared from Valonia. Consequently, these assays should be treated as a relative and not quantitative assessment. The nature of the original substrate selected—especially its extent of crystallinity—should always be taken into account. Proper controls and reference substrates should always be used. One should be wary about comparison among results reported by different laboratories and even by different researchers in the same laboratory. Nevertheless, such assays give an excellent indication of whether a given enzyme preparation exhibits substantial activity toward crystalline cellulose substrates.

Endoglucanase Versus Exoglucanase Activity As discussed earlier in this chapter, the cellulases have traditionally been divided into either endoglucanases or exoglucanases (> Fig. 6.4). The biochemical or enzymatic assays that discriminate between these two modes of action usually involve soluble forms of cellulose, i.e., carboxymethyl or hydroxymethyl derivatives of cellulose. The action of a given enzyme on these substrates is followed by determining the amount of reducing ends generated by the enzyme and the degree of polymerization (DP). The reducing power is usually determined either by using reagents such as 3,5-dinitrosalicylic acid (DNS) (Miller et al. 1960), ferricyanide (Kidby and Davidson 1973), or copperarseno molybdate (Green et al. 1989; Marais et al. 1966). Despite their traditional popularity, these two methods are intrinsically disadvantageous, owing to interference by metal ions and certain buffers. Moreover, such assays are sensitive to the chain length of the reducing end. A more recent approach involves the use of disodium 2,20 -bicinchoninate (BCA) for determination of reducing sugar. This procedure is more sensitive than the conventional methods and gives comparable values of reducing sugars for cellodextrins of different lengths (Doner and Irwin 1992; Garcia et al. 1993; Vlasenko et al. 1998; Waffenschmidt and Jaenicke 1987). Viscosity-based measurements represent the most common approach for assessing the degree of polymerization. This approach is highly sensitive for internal bond cleavage, which leads to significant reduction of the average molecular weight of the substrate. The comparison between the amount of reducing sugars generated and the average molecular weight

(i.e., viscosity or fluidity of the soluble cellulose substrate) gives a very good indication whether an enzyme is essentially exo- or endo-acting. The average degree of polymerization can also be evaluated by size-exclusion chromatography either alone (Srisodsuk et al. 1998; Teeri 1997) or combined with multi-angle laser light scattering (Vlasenko et al. 1998). Mass spectrometric procedures can also be applied to determine the identity and distribution of degradation products following hydrolysis of cellulosic substrates by an enzyme (Hurlbert and Preston III 2001; Rydlund and Dahlman 1997). The mode of enzymatic action can also be appraised by determining the increase in reducing power associated with the insoluble versus the soluble fraction of the substrate. Increase in the proportion of reducing sugars associated with the soluble fraction indicates an exo type of activity whereas a relatively large increase in the insoluble fraction would suggest an endo type of activity (Barr et al. 1996). Exocellulases can exhibit different specificities depending on their preference for the reducing or nonreducing end of the cellulose chain (Barr et al. 1996; Teeri 1997). This feature of an exocellulase can be determined either by using oligosaccharide substrates labeled by tritium or 18O at the reducing end. Other procedures involve NMR, HPLC, and/or mass spectrometric analysis of products released from native (unlabeled) cellooligosaccharides. In previous studies, the 3D structures of enzymesubstrate complexes have been obtained, and the specificities of the enzyme can be interpreted directly from the data (Davies and Henrissat 1995; Davies et al. 1998; Divne et al. 1998; Juy et al. 1992; Notenboom et al. 1998; Parsiegla et al. 1998; Rouvinen et al. 1990; Sakon et al. 1997; Zou et al. 1999). These efforts have since continued as novel families of the glycoside hydrolases were established; selected members of these families were subjected to crystallization studies in order to characterize the overall structural features and mode of action of the entire family.

Processivity One of the major recent conceptual advances in assessing the mode of enzymatic action of a cellulase is the concept of processivity. Processive enzyme action can be defined as the sequential cleavage of a cellulose chain by an enzyme. In effect, exoglucanases are by nature and structure processive enzymes. Their tunnel-like active site thus allows processive action on the cellulose chain. Endoglucanases, however, were thought to be intrinsically non-processive. However, the traditional distinction between exo- and endo-cellulases has been modified. Experiments combining two or more purified cellulases have shown that synergism can even be detected upon mixing two different types of exo-acting enzymes. Such experiments led to the recognition that the exo enzymes can operate on both ends (i.e., the reducing and nonreducing ends) of the cellulose chain. Some enzymes, however, exhibit both endo and exo activities, although in such cases the endo-cellulase activity is usually very low. In attempts to explain these phenomena, the concept of

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

processivity was proposed, by which the activity of the enzyme is characterized by the sequential hydrolysis of the cellulose chain. Implicit in this concept is the notion that the catalytic site of the enzyme remains in continual and intimate contact with a given chain of the cellulose substrate. A more complete mechanistic picture of the processive nature of such cellulases was revealed with the advent of highresolution 3D structures. It was thus demonstrated that the cellulose chain makes contact with the protein at multiple sites, either via a tunnel-shaped structural element (such as that observed in the Family-48 enzymes) or by a special type of CBM (such as the Family-9 Theme B cellulases). These arrangements allow the threading of the cellulose chain into the active site, and, following initial cleavage at the end of the chain, the enzyme can move along the chain and position itself for the next cleavage. In addition to this processive nature of the active site, these enzymes can also make classic endo cleavages thus generating new ends. Biochemically processive enzymes exhibit characteristics between endo and exo enzymes. They have low but detectable endo activity toward soluble derivatives of cellulose (i.e., CMC), and may or may not possess exo activity on such substrates. With insoluble substrates, they will generate reducing power with a ratio between the soluble to the insoluble fractions of about 7. Endocellulases usually give a ratio of less than 2, whereas exocellulases produce a ratio of 12–23 (Irwin et al. 1998). Once the processive nature of an enzyme has been indicated experimentally, molecular insight into the mechanisms responsible for this feature can be gained by determining the 3D crystal structure of the active site together with model cellodextrins. In the case of the cellulases, the crystal structure of the catalytic module together with the fused CBM, combined with accumulating enzymatic activity data, allowed further postulation as to the accessory role of the fused module. The fused CBM presumably interacts with a single cellulose chain and feeds it into the active site. Interestingly, this module does not bind crystalline cellulose, but is inferred to act in dynamic binding of the single cellulose chain prior to its hydrolysis, thereby imparting the quality of processivity to the enzyme. Once such a property is associated with a given type of enzyme, the primary structure of the protein can now be used as an indication for all such enzymes. In the case of the Family-9 Theme B enzymes, it is now possible to identify the catalytic module (e.g., glycoside hydrolase Family 9) and the additional accessory modules (in this case, Family-3c CBM). Thus, the primary structure may by itself give a strong indication of the nature of the enzyme itself. Of course, the ultimate identification as to the mechanism of enzyme activity will come from detailed 3-D structure of the enzyme-substrate complex. An intriguing recent development in the analysis of the cellulolytic action of a given cellulase or a mixture of cellulase is the direct transmission electron microscopic (TEM) observation of the enzymatic action on bacterial cellulose ribbons. The approach provides information as to the endo or exo preference of the enzyme, the extent of processivity, as well as the

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directionality of hydrolysis (i.e., from the reducing to the nonreducing ends or vice versa). This strategy has been used to study the hydrolysis of bacterial cellulose ribbons by individual purified enzymes, mixtures of purified enzymes, and intact cellulosomes.

Mechanism of Catalysis The mechanism of catalysis of cellulases address issues such as stereochemistry, binding- and active-site residues, and transition state intermediates. Excellent reviews have been published covering many of these issues (Ly and Withers 1999; McCarter and Withers 1994; Rye and Withers 2000; Sinnott 1990; White and Rose 1997; Withers 2001; Withers and Aebersold 1995; Zechel and Withers 2000). The fact that the stereochemistry and catalytic residues are conserved between members of the same family allows the putative identification of these elements if one member of the given (glycoside hydrolase) has been characterized biochemically (Henrissat and Bairoch 1996; Henrissat et al. 1995; Henrissat and Davies 1997). The stereochemistry of the reaction can in most cases be determined by proton NMR spectroscopy or by using chromatography systems that allow the resolution of anomeric species. In the case of NMR, the reaction between the test enzyme and its substrate is carried out in D2O and the appearance of the anomeric proton can be easily detected. Thus, for the degradation of cellulose, a retaining enzyme would produce a product in the b configuration whereas an inverting enzyme would yield the a-sugar. The catalytic residues can be identified by performing sitedirected mutagenesis on conserved acidic residues and studying the catalytic properties of the mutants with substrates bearing different leaving groups. Commonly used phenol substituents include the following, listed in order of leaving group ability (pKa values shown parenthetically): 2,4-dinitro (3.96) > 2,5dinitro (5.15) > 3,4-dinitro (5.36) > 2-chloro4-nitro (5.45) > 4-nitro (7.18) > 2-nitro (7.22) > 3,5-dichloro (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. 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 kinetics-based 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

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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 for substrates with a 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 characterized by free enzymes, cell-bound enzymes, multifunctional enzymes, cellulosomes, or any combination of the latter. Collectively, these four types of enzymes represent the major paradigms of plant cell wall polysaccharide-degrading enzymes (Himmel et al. 2010). 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) 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 modules. Such enzymes often specialize on 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 module 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 module on the substrate. The intact free enzyme, however, remains unattached to other enzymes and can work in an independent manner on a given substrate. Free enzymes containing larger numbers of ancillary modules are also prevalent in components of bacterial cellulase systems. Examples of bacteria that possess free carbohydratedegrading enzymes include the well-established actinomyces, Thermobifida fusca and Cellulomonas fimi. More details of their enzyme systems will be presented in a forthcoming section. The more recent discovery of Saccharophagus degradans 2-40 has provided a particularly intriguing and elaborate cellulolytic bacterium that can grow alone on cellulose without the assistance of other microorganisms. S. degradans 2-40 is the first freeliving marine bacterium demonstrated to be capable of degrading cellulosic algae and higher plant material, and its genome codes 15 extraordinarily long polypeptides, ranging from 274 to 1,600 kDa (Weiner et al. 2008). This bacterium has a remarkable range of catabolic capabilities, and many of the enzymes exhibit unusual modular architectures including novel combinations of catalytic and substrate-binding modules. S. degradans 2-40 can degrade different complex polysaccharides (at least 10), including agar, chitin, alginic acid, cellulose, b-glucan, laminarine, pectin, pullulan, starch, and xylan and utilize them as sole carbon and energy sources (Ensor et al. 1999). The genome of S. degradans encodes abundant glycoside hydrolases families mainly GH5 (20 in number) followed by GH43 (13), GH13 (10), GH16 (9), GH2 (7), and GH3 (6)

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

(Taylor et al. 2006). The CAZymes of this bacterium are generally extracellular free enzymes, many of which are decorated with at least one CBM. One of the GH5 enzymes is believed to exhibit processive endoglucanase activity and contains two such catalytic modules together with three copies of family-6 CBMs in the same polypeptide chain (Watson et al. 2009). Interestingly, the chitinases, agarases, and alginases produced by this bacterium are not exported into the extracellular matrix but are localized in surface protuberances, resembling those of the cellulosomeproducing bacteria. The genome of S. degradans encodes the largest set of identifiable CBMs so far reported (Weiner et al. 2008). Carbohydrate binding modules of Family 6 (CBM6) are the most numerous (43 copies) followed by CBM 32 (26) and CBM2 (19). Among the long polypeptides encoded by S. degradans 2-40 genome, five of them contain at least 52 bacterial cadherin (CA) and cadherin-like (CADG) domains. Both domain types exhibit Ca2+-dependent binding to different complex polysaccharides which serve as growth substrates (Fraiberg et al. 2010, 2011). Recent evidence suggests that the regulatory mechanisms that control the expression of the various enzymes of the cellulolytic system are very complex and contain an intricate chemotaxis signal transduction network for detecting both extracellular and intracellular signals and numerous chemotactic response regulators (Zhang and Hutcheson 2011).

Multifunctional Enzymes Some cellulases exhibit a more complex architecture in that more than one catalytic module and/or CBM may be included in the same protein. Examples of such enzymes are the very

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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 module. Additional examples of the latter type of multifunctional enzyme have been found in A. cellulolyticus and C. clariflavum. Other paired catalytic modules include those from Family 44 and either Family 5 or 9. Such an arrangement might indicate a close cooperation between two particular catalytic modules, which may lead to synergistic action on the cellulosic substrate, thus portending on a smaller scale the advent of cellulosomes. Like the cellulases, xylanases also tend to exhibit a modular structure, being composed of multiple modules joined by linker sequences. Family-10 and Family-11 xylanases may be linked in the same polypeptide chain either to each other, to catalytic modules 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. 6.3). In this manner, the xylan chain would be severed by the xylanase component (Xyn in > Fig. 6.3) and the lignin-xylan association would be disconnected simultaneously by the feruloyl acid esterase (Fae in > Fig. 6.3). Indeed, some xylanases are extremely complex in their modular architecture (> Fig. 6.10). 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 cellulosespecific CBM? Unlike the case of various cellulases, for which the CBM is usually essential for degrading insoluble crystalline

. Fig. 6.10 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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cellulose, the CBMs of a hemicellulase do not necessarily bind the hemicellulose component (xylan). In some cases, its CBM is in fact an authentic CBM 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 CBMs (CBM3) shown in > Fig. 6.10 apparently bind to crystalline cellulose. Why would this xylanase require three tandem copies of the same type of CBM is yet another mystery that should eventually be addressed experimentally. At any rate, once bound via the 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 in order to direct the appropriate catalytic module to its true substrate. Hence, the modular proximity of the xylanase shown in > Fig. 6.10 would presumably indicate that the two CBM22 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. 1983a, b). Since its initial description in the literature, the cellulosome concept has been subject to numerous reviews (Bayer et al. 1996; Be´guin 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 1988a, b, 1991, 1993; Lamed et al. 1983; Shoham et al. 1999). Cellulosomes in C. thermocellum exist in both cell-associated and extracellular forms, the cell-associated form being associated with polycellulosomal protuberance-like organelles on the cell surface. Later, cellulosomes were detected in other cellulolytic organisms (Lamed et al. 1987a, b; 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. 6.11). The role of surface functions was further shown to be important in increasing the efficiency of cellulose fermentation (Lu et al. 2006). 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

. Fig. 6.11 A very large, multi-modular xylanase from Caldicellulosiruptor. The 1795-residue enzyme contains 8 separate modules, including 2 catalytic modules from Family-10 (invariably a xylanase) and Family-43 (frequently an arabinofuranosidase). These are modulated by numerous CBMs, which include three from Family-3 (likely for binding to crystalline cellulose), two from Family-22 (shown to function in xylan binding) and one from Family-6

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 multi-subunit nature of the cellulosome (> Fig. 6.12). 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. These efforts were further extended with the genome sequences of various Clostridia and Ruminococci species.

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 bacteria 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 Family-5 cellulase and a Family-13 amylase-pullulanase from Bacillus, a Family-10 xylanase from Caldicellulosiruptor (Saul et al. 1990), a Family-5 endoglucanase from Clostridium josui, a Family-16 lichenase and a 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 amylase-pullulanase) from different species of Thermoanaerobacter (Matuschek et al. 1996). The

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

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. Fig. 6.12 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, b). Without pretreatment with cationized ferritin, these structures are invisible. In (b), the cellulosebound 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, notably C. thermocellum (Bayer and Lamed 1986)

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. 6.13). Other surface functions, such as adhesive properties, may also be associated with the same protein (Fraiberg et al. 2011; Ozdemir et al. 2012). In a different bacterium, Ruminococcus flavefaciens, from the rumen of herbivores, the cellulosome is attached covalently to the bacterial cell surface by scaffoldin E (ScaE) (Rincon et al. 2005). ScaE is an anchoring scaffoldin that includes a C-terminal cell-anchoring signal motif for covalent attachment to the cell wall via the enzymatic action of an appropriate cell-associated sortase. Two key scaffoldins, scaffoldin B (ScaB) and cellulosebinding protein A (CttA), are attached to the cell-surface ScaE scaffoldin via a C-terminal X-dockerin (XDoc) modular dyad. ScaB is essentially an adaptor scaffoldin to which scaffoldin A (ScaA) and/or selected dockerin-bearing enzymes, including cellulases, are incorporated into the R. flavefaciens cellulosome. CttA is believed to mediate the attachment of the bacterium to cellulosic substrates (Rincon et al. 2007). Genome sequencing revealed several other structural proteins that include sortase signal motifs (Berg Miller et al. 2009; Rincon et al. 2010). In one case, a GH10 xylanase was sequenced that also bears a sortase signal motif at its C terminus. An additional mechanism of bacterial surface attachment has recently been reported (Devillard et al. 2004; Xu et al. 2004; Ezer et al. 2008). Several CBMs were discovered following genomic sequencing of the rumen bacterium, R. albus. These CBMs were classified as Family-37 and found exclusively in R. albus. Half of the parent proteins are carbohydrate-acting

enzymes (glycoside hydrolases, pectate lyases, and carbohydrate esterases). The involvement of CBMs in anchoring plant cell wall–degrading enzymes onto the bacterial cell surface extends the types of functions that this superfamily of protein modules performs in nature.

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. 6.14. The cellulosomal enzyme subunits are united into a complex by means of the primary scaffoldin subunit (Bayer et al. 1994; Shoseyov et al. 1992; Fujino et al. 1993; Gerngross et al. 1993). The scaffoldins usually contain a Family-3 CBM that provides the cellulose-binding function (Poole et al. 1992). The scaffoldins also contain multiple copies of a definitive type of cohesin module. The cellulosomal enzyme subunits, on the other hand, contain a complementary type of dockerin module. The interaction between the cohesin and dockerin modules provides the definitive molecular mechanism that integrates the enzyme subunits into the cellulosome complex (Salamitou et al. 1994b; Tokatlidis et al. 1991, 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. 1998a). This has indeed been confirmed in many cases. However, noncellulosomal cohesins and dockerins have been identified in many bacteria, as well as archaea and a few isolated cases of primitive eukarya, without a link to polysaccharide degradation

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. Fig. 6.13 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))

(Bayer et al. 1999; Adams et al. 2008; Chitayat et al. 2008a, b; Peer et al. 2009; Voronov-Goldman et al. 2009). The major difference between free enzymes and cellulosomal enzymes is that the free enzymes usually contain a CBM for guiding the catalytic module to the substrate, whereas the cellulosomal enzymes carry a dockerin module that incorporates the enzyme into the cellulosome complex. Otherwise, both the free and cellulosomal enzymes contain very similar types of catalytic modules. The cellulosomal enzymes rely on the Family-3a CBM 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 nine-stranded beta sandwich with a jelly-roll topology (Shimon et al. 1997; Spinelli et al. 2000; Tavares et al. 1997).

The dockerin module 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 EF-hand motif of various calcium-binding proteins (e.g., calmodulin and troponin C). Within this repeated sequence is a 12-residue calcium-binding loop, indicating that calciumbinding is an important characteristic of the dockerin module. This assumption was eventually confirmed experimentally (Yaron et al. 1995). The specificity characteristics of the cohesin-dockerin interaction have also been investigated. The results showed that four suspected residues may serve as recognition codes for interaction with the cohesin module (Mechaly et al. 2000, 2001; Page`s et al. 1997). The three-dimensional solution structure of the 69-residue dockerin module of a Clostridium thermocellum cellulosomal cellulase subunit was determined (Lytle et al. 2001). As predicted earlier (Bayer et al. 1998; Lytle et al. 2000; Page`s 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 module.

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

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. Fig. 6.14 Simplified schematic view of the molecular disposition of the cellulosome and one of the associated anchoring scaffoldins 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 scaffoldin to cellulosome to cellulose substrate is illustrated. The SLH module links the parent anchoring scaffoldin to the cell. The cellulosomal scaffoldin subunit performs three separate functions, each mediated by its resident functional modules: (1) its multiple type-I cohesins integrate the cellulosomal enzymes into the complex via their resident type-I dockerins, (2) its Family-3a CBM binds to the cellulose surface, and (3) its type-II dockerin interacts with the type-II cohesin of the exocellular anchoring scaffoldin

The scaffoldin of C. thermocellum also contains a special type of dockerin module. This dockerin failed to bind to the cohesins from the same scaffoldin subunit, but instead interacted with a different type of cohesin—termed ‘‘type-II’’ cohesins—identified on the basis of sequence homology (Salamitou et al. 1994a). These cohesins are somewhat different from those of type I, having an additional segment and diversity in the latter half of the sequence. Three-dimensional structures for several examples of type-II cohesins have been reported (Noach et al. 2003, 2005, 2008, 2009; 2010; Carvalho et al. 2005). The type-II cohesins were discovered as component parts of a group of noncatalytic cell-surface ‘‘anchoring’’ proteins on C. thermocellum (Leibovitz and Be´guin 1996; Leibovitz et al. 1997; Lemaire et al. 1995; Salamitou et al. 1994a). The three known anchoring scaffoldins in C. thermocellum contain different copy-numbers of the typeII cohesins as illustrated in > Fig. 6.15. Each of these anchoring scaffoldins also contains an SLH (S-layer homology) module, analogous to those of the cell-bound enzymes mentioned above (see section on > ‘‘Cell-Bound Enzymes’’). The intervening sequences, however, between the cohesins and SLH modules are different. In any case, the type-II 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. In recent years, structures for cohesin-dockerin complexes have been reported, which represent a significant breakthrough in our understanding of how the scaffoldins are organized and cellulosome architecture in general. In this context, cohesindockerin complexes for both type I and type II have been elucidated. Moreover, the structures provide insight on the molecular level regarding the specificity of this high-affinity interaction. The crystal structure of C. thermocellum scaffoldin-borne cohesin two module together with the dockerin module from xylanase 10B was the first cohesin-dockerin complex reported (Carvalho et al. 2003). Interestingly, very little conformational change was observed in the cohesin module relative to the known structure of the same cohesin alone. The dockerin bound to the 8-3-6-5 face of the cohesin via an extensive hydrogen-bonding network and supporting hydrophobic interactions. Surprisingly, the twofold symmetry observed for the type-I dockerin sequences of this bacterium reflected a 180 rotation on cohesin surface, resulting in a dual mode of binding, in which the parent enzyme can attain one of two

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very different conformations in space, with respect to the interacting modular couterparts (Carvalho et al. 2007; Pinheiro et al. 2008). The subsequent crystal structure of the type-II complex between the C. thermocellum SdbA cohesin module and the CipA scaffoldin XDoc modular dyad provided additional surprises (Adams et al. 2005). The resultant complex structure exhibited striking differences from that of the type-I complex. Notably, the lack of sequential symmetry of the dockerin module appeared to preclude a dual mode of binding. Indeed, as opposed to the type-I cohesin-dockerin interaction, the type-II

dockerin contacts the cohesin counterpart across the entire length of both helices, which appears to result in a higher affinity and a single mode of interaction.

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. 6.15. Two scaffoldins for Acetivibrio cellulolyticus and

. Fig. 6.15 Schematic view of the modular similarity and diversity of scaffoldins from different cellulosome species. Four major scaffoldins of the current C. thermocellum paradigm are shown. The type-I cohesin-dockerin pairs are shown in yellow, the type-II pairs are shown in pink, and the anchoring component (the SLH module) is in green. Anchoring scaffoldins are designated by the adjacent symbol of an anchor. Other mesophilic clostridial species are characterized by a single scaffoldin. The four scaffoldins of the A. cellulolyticus system are more cross-interactive than that of the C. thermocellum paradigm. The reversed types of cohesin-dockerin pairings are evident in the B. cellulosolvens system, as are its two exceptionally large scaffoldins. The type-III cohesin-bearing scaffoldins of the R. flavefaciens system are especially elaborate. The single-cohesin ScaC ‘‘adaptor’’ scaffoldin provides the means with which to modify the repertoire of cellulosomal components, and the monovalent ScaE cohesin attaches the ScaB adaptor scaffoldin to the cell surface. Each of the seven ScaB cohesins binds to a cohesin of the trivalent ScaA primary scaffoldin which incorporates dockerin-bearing enzymes into the complex. Micrographs of the different bacteria are included in the figure

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

Bacteroides cellulosolvens, like C. thermocellum, carry dockerin modules at their C terminus (Ding et al. 1999, 2000). The A. cellulolyticus genome also includes a gene (immediately downstream of the scaffoldin gene) coding for an anchoring scaffoldin, that contains type-II cohesins. 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 scaffoldins. The scaffoldins from the other clostridial species thus far described all lack ‘‘type-II’’ dockerin modules, the inference being that cells of C. cellulovorans, e.g., would apparently not bear anchoring scaffoldins which contain type-II cohesins. Since their cellulosomes appear to be surface bound, their anchoring thereto is likely accomplished via an alternative molecular mechanism. In subsequent publications (Doi and Tamura 2001; Tamaru and Doi 1999; Tamaru et al. 1999), a cell-surface binding function was proposed for a module of unknown function [designated X2 (Coutinho and Henrissat 1999a, b, c)] 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 module may be recognized as an SLH module. The four X2 modules of the C. cellulovorans scaffoldin are very similar in sequence to the Xmodules from the scaffoldins of Clostridium cellulolyticum and C. josui, which contain only two and one copy of this module, respectively. If this module 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 module 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 1999a, b, c)] that the authors consider to be involved in cell-surface attachment (Tamaru and Doi 1999). In any case, final proof of the function of the X2 and X48 modules awaits biochemical examination, as has been clearly achieved for the SLH module of the C. thermocellum anchoring scaffoldins (Chauvaux et al. 1999; Lemaire et al. 1998). Finally, two novel scaffoldins were sequenced from the rumen bacterium, Ruminococcus flavefaciens strain 17 (Ding et al. 2001; Rincon et al. 2003, 2004, 2007). Although the proteins contain 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 CBM. The lack of a scaffoldin CBM and the question as to how the ruminococcal cellulosome(s) and/or the bacterium bind to the substrate were eventually resolved at least partially by the discovery of an additional CBM-bearing scaffoldin coded by a gene in the scaffoldin gene cluster of this bacterium (Rincon et al. 2007). Furthermore, a draft genome sequence of a similar strain of the same species was recently reported (Berg Miller et al. 2009), which revealed an

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exceptionally elaborate cellulosome system with a multitude of dockerin-bearing components (Rincon et al. 2010), roughly threefold of that observed in the C. thermocellum genome.

Schematic Comparison of Prokaryotic Cellulase Systems In this section, we will describe schematically the similarity of and diversity in representative enzyme systems, demonstrating different strategies, from 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 with 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. Indeed, during the past decade, the genomes of many cellulolytic species have been sequenced (see > Table 6.1), thereby supplementing our knowledge of the cellulase and cellulosome components. Representative schematic lists of the latter components will be provided below in forthcoming figures. More extensive descriptions of the total content of carbohydrate-active enymes, i.e., the CAZome, of the different cellulolytic bacteria, are now readily obtainable via the CAZy database (http://www.cazy.org/). 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 glycoside hydrolases, including the different cellulases, per se; the hemicellulases (e.g., xylanases, mannanases); pectin-degrading enzymes; etc. The cellulase system of the mesophilic cellulolytic aerobe, Cellulomonas fimi, is one of the first studied, and for many years has 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. The modular composition and family associations of representative glycoside hydrolases from this

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. Fig. 6.16 Cellulomonas fimi cellulase system – an example of a cell-free enzyme system: Pictographic view of the enzyme components and their modular architecture. 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 modules are enumerated; the catalytic modules are in red. Key to symbols: GH glycoside hydrolase (e.g., cellulase, xylanase, mannanase), CE carbohydrate esterase (e.g., acetyl xylan esterase and ferulic acid esterase), CBM carbohydrate-binding module), SLH S-layer homology (module), FN3 fibronectin-3 (domain), Ig immunoglobulin-like domain, X domain of unknown function

bacterium are shown symbolically in > Fig. 6.16. As an example of a free enzyme system, most of the enzymes bear a substratetargeting CBM, which, in Cellulomonas fimi, are mainly from Family-2. Several of the enzymes have multiple copies of the FN3 domain (fibronectin 3 domain), the function of which is still unknown. The Cellulomonas system includes four Family-6 enzymes. Two of these are shown in the figure—an endoglucanase and an exoglucanase (cellobiohydrolase) of the types described in > Fig. 6.4. The modularity of the endoglucanase is very simple, having the Family-6 catalytic module together with a Family-2 CBM. The cellobiohydrolase is a bit more complex with 3 additional FN3 domains that separate the same two types of modules. The two additional Cel6 enzymes appear to lack CBMs and are not included in the figure. 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. 6.7). Two additional Family-9 cellulases are included in Cellulomonas fimi; one contains a simple GH9 catalytic module

with a single CBM2 and the other has no additional ancillary modules (neither are described in the figure). In addition, a simple Family-5 cellulase and an interesting cell-borne Family-26 mannanase are components of the system. An additional Family-5 enzyme bears a CBM13 and two other Family-26 enzymes are present (not shown). The fact that an enzyme, i.e., the Family-26 enzyme, bears an SLH module and is presumably cell-associated would underscore its importance to the cell. Finally, 3 xylanases are part of the enzymatic apparatus of Cellulomonas fimi. One of these xylanases is a simple enzyme consisting of a Family-10 catalytic module connected to a Family-2 CBM. The other two are more complicated, each containing two catalytic modules—either a Family-10 or a Family-11 module and a carbohydrate esterase (in both cases, probably an acetyl xylan esterase (> Fig. 6.3)—plus several CBMs. The genome for this bacterium has recently been sequenced, and its enzymatic system is much more extensive than that shown in the figure. For example, seven members of GH43 have been detected in its genome. For more information regarding the CAZome of Cellulomonas fimi, the reader is referred to the CAZy database (http://www.cazy.org/). A second example of a free enzyme system, from the aerobic thermophilic bacterium Thermobifida fusca (formerly classified

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

6

. Fig. 6.17 Thermobifida fusca cellulase system. A cell-free enzyme system. Compare with the Cellulomonas system (> Fig. 6.16). Key to symbols: GH glycoside hydrolase (e.g., cellulase, xylanase, mannanase), CBM carbohydrate-binding module, FN3 fibronectin-3 (domain), Ig immunoglobulin-like domain, X domain of unknown function

as Thermomonospora fusca), has also been studied extensively (Wilson 1992, 2004, 2008, 2009; Wilson and Irwin 1999). A brief comparison of its known enzyme components (> Fig. 6.17) shows a striking resemblance to those of Cellulomonas (compare > Figs. 6.16 and > 6.17). Both species produce similar types of cellulases from families 5, 6, 9, and 48 plus xylanases from families10 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 module. 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 T. fusca genome has now been sequenced, and more extensive information is available regarding its CAZome (http://www.cazy.org/). In contrast to the numerous members of the GH43 enzymes in Cellulomonas fimi, there is only one GH43 enzyme in T. fusca. 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, Fibrobacter succinogenes, Saccharophagus degradans, and various species of Streptomyces, Erwinia, and Thermatoga, appear to have several cellulases, xylanases, and mannanases from the common families, together with other glycoside 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 cellulose-binding module, 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 modules of unknown function (X modules). Once again, until the genome sequences of cellulolytic prokaryotes are widely available, we are still limited in our capacity to compare among the enzyme systems, due to our incomplete knowledge of their enzyme sequences.

Multifunctional Enzyme Systems In an hyperthermophilic bacterium, 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. 6.18). As mentioned earlier (see section > ‘‘Multifunctional Enzymes’’), the appearance of two catalytic modules in the same enzyme would infer a distinctive synergistic action between the two. Thus, in Caldicellulosiruptor CelA, the Family-9 and Family-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

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. Fig. 6.18 Caldicellulosiruptor enzyme system: An example of a cell-free enzyme system that includes several multifunctional enzymes. Key to symbols: GH glycoside hydrolase (e.g., cellulase, xylanase, mannanase), CBM carbohydrate-binding module, SLH S-layer homology (module). See also > Table 6.5

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 an SLH module (Ozdemir et al. 2012). 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 enzymes is usually one or more copies of a Family-3 CBM. The presence of more than one copy of a CBM in this case may reflect the extreme temperatures of the ecosystem. Other bacterial strains that include at least one free bifunctional enzyme in their enzyme systems are Anaerocellum thermophilum (now considered a species of Caldicellulosiruptor), Bacillus stearothermophilus, Fibrobacter succinogenes, Prevotella ruminicola, Ruminococcus albus, Ruminococcus flavefaciens, Streptomyces chattanoogensis, and thermophilic anaerobe NA10. The genomes of several species of Caldicellulosiruptor have now been sequenced (Kataeva et al. 2009; Blumer-Schuette et al. 2011). Each is characterized by different sets of bifunctional enzymes (> Table 6.5), and some of these genomes either lack gene coding for such enzymes altogether or contain only one or two. Others carry up to seven bifunctional enzymes in their respective genomes (Dam et al. 2011). The different bifunctional

enzymes include the various combinations (Himmel et al. 2010), notably cellulase-cellulase, cellulase-hemicellulase, hemicellulasehemicellulase, hemicellulase-carbohydrate esterase, and even polysaccharide lyase-hemicellulase forms. The multiplicity of these genomes indicates the diverse nature of this genus of hyperthermophilic bacteria and reflects different patterns of substrate utilization.

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 (see earlier section > ‘‘Similarity and Diversity of Scaffoldins from Different Species’’), in the case of Clostridium thermocellum, Acetivibrio cellulolyticus, Bacteroides cellulosolvens, and Ruminococcus flavefaciens, the cellulosomes appear to be attached to the cell surface. The cellulosomes of C. cellulolyticum, C. cellulovorans, and C. josui may also be cell-associated, but, if so, the lack of a scaffoldin-borne dockerin and reciprocal anchoring scaffoldin would suggest an alternative mechanism. The cellulosomes of some mesophilic clostridia, such as C. cellulolyticum, C. cellulovorans, C. josui, and C. papyroslvens

6

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Table 6.5 Bifunctional enzymes from the genus Caldicellulosiruptor Caldicellulosiruptor speciesa Enzyme (modular components)

Athe

GH5-CBM3-CBM3-CBM3-GH5

+

+ +

GH5-CBM3-CBM3-GH44

+

GH9-CBM3-CBM3-CBM3-GH5

+

GH9-CBM3-CBM3-CBM3-GH48

+

Calhy

CBM22-CBM22-GH10-CBM9-CBM9-CBM9-CE15

Calkr

Calkro

Calla

COB47

Calow

+ +

+

+

+

+

GH10-CBM3-GH5

+

GH10-CBM3-CBM3-GH5 GH10-CBM3-CBM3-GH48

+ +

+

CBM54-GH16-GH55-CBM32 GH43-CBM22-GH43-CBM6

+ +

GH74-CBM3-CBM3-GH44 GH74-CBM3-CBM3-GH48

Csac

+

+

+

+ +

PL11-CBM3-CBM3-CBM3-GH44

+

+

+

+

a

Athe, Anaerocellum thermophilum DSM 6725 (Caldicellulosiruptor bescii); Calhy, Caldicellulosiruptor hydrothermalis 108; Calkr, Caldicellulosiruptor kristjanssonii 177R1B; Calkro, Caldicellulosiruptor kronotskyensis 2002; Calla, Caldicellulosiruptor lactoaceticus 6A; COB47, Caldicellulosiruptor obsidiansis OB47; Calow, Caldicellulosiruptor owensensis OL; Csac, Caldicellulosiruptor saccharolyticus DSM 8903

are very similar. The genes encoding for many or most of the enzymes in the latter cellulosomal systems are arranged in a large cluster on the chromosome (> Fig. 6.19). Additional 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. 2000). In contrast, the cellulosomal genes from C. thermocellum are generally scattered over a large portion of the chromosome (Guglielmi and Be´guin 1998). A few small clusters of cellulosomal genes are apparent in the genome, including a scaffoldin-containing cluster (> Fig. 6.19) 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 microorgansims. The genomes of all three bacteria have been sequenced, and the genomes of other cellulosome-producing bacteria are forthcoming in the near future. 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 for this species contains more than one catalytic module (> Fig. 6.20). The largest one, Cel9E (estimated at 94 kDa), is a Theme-D Family-9 cellulase (Gaudin et al. 2000). The critical Family-48 cellulase (Cel48F) is also a major cellulosome component (Reverbel-Leroy et al. 1997). The gene cluster of C. cellulolyticum contains several copies of other Family-9 cellulases, including Cel9G, Cel9H, and Cel9J, all of which contain the Theme-B fused Family-3c CBM (Belaich et al. 1998) (> Fig. 6.8). The cellulosome system in this

bacterium also contains numerous Family-5 cellulases (including Cel5A and Cel5D), a Family-5 mannanase (Man5K, which bears an N-terminal rather than C-terminal dockerin) and a Family-8 cellulase (Cel8C). Biochemical characterization of the C. cellulolyticum cellulosome demonstrated by SDS-PAGE a 160-kDa scaffoldin band and up to 16 smaller bands, representing putative enzyme subunits (Gal et al. 1997b). Many of these were clearly identified as known gene products. Early biochemical evidence suggested that xylanases from C. cellulolyticum are also organized in a cellulosome-like complex (Mohand-Oussaid et al. 1999). The genome sequence of this bacterium and subsequent proteomics studies revealed 62 dockerin-containing proteins, most of which are enzymes, including cellulases, xylanases, and other glycoside hydrolases, as well as carbohydrate esterases and polysaccharide lyases (Desvaux 2005; Blouzard et al. 2010). Cellulosomal Components from Clostridium cellulovorans

Like C. cellulolyticum, the cellulases from this organism are relatively simple (see pictographical description of representative enzymes in > Fig. 6.21). In addition to the cellulosomal enzymes thus described, several non-cellulosomal endoglucanases have also been partially or totally sequenced (Doi et al. 1998; Tamaru et al. 1999), notably those from Family-9 (Kosugi et al. 2002; Han et al. 2004, 2005). Several of the cellulosomal enzymes are architecturally synonymous to those of the C. cellulolyticum system (compare > Figs. 6.20 and > 6.21). This includes the critical Family-48 cellulase (Exg48S) (Liu and Doi 1998), two copies of the ThemeB Family-9 cellulase (Eng9H and Eng9Y), a Family-5 endoglucanase, and a Family-5 mannanase that bears an

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Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Fig. 6.19 Cellulosome-related gene clusters. Enzyme-linked gene clusters of the mesophilic Clostridia include an initial primary scaffoldin gene followed downstream by a series of genes encoding for various dockerin-bearing enzymes. Note the extensive similarity and subtle differences in the succession of enzyme-encoding genes. Multiple-scaffoldin gene clusters of the indicated bacteria comprise two or more genes in tandem that encode for scaffoldins. Scaffoldin genes and genes for enzymes are shown as light blue and pink arrows, respectively, whose length gives the approximate proportional size of the given gene relative to the others

N-terminal dockerin (Tamaru and Doi 2000). Rather than a single Theme-D Family-9 cellulase as in C. cellulolyticum, the C. cellulovorans system contains two such enzymes (Eng9K and Eng9M). The C. cellulovorans cellulosome also appears to contain an unusual Theme-A Family-9 cellulase (Eng9L) that lacks helper modules. 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. The genome of C. cellulovorans was sequenced recently (Tamaru et al. 2010). Interestingly, 57 cellulosomal genes were identified in the genome, which, in addition to carbohydrateactive enzymes, also coded for lipases, peptidases, and proteinase inhibitors. Cellulosomal Components from Clostridium thermocellum

Compared to the cellulosomal systems of C. cellulovorans and C. cellulolyticum, the enzymes from C. thermocellum are

relatively large proteins, ranging in molecular size from about 40 to 180 kDa (Bayer et al. 1998c, 2000; Be´guin and Lemaire 1996; Felix and Ljungdahl 1993; Lamed and Bayer 1988; Shoham et al. 1999). Examination of > Fig. 6.22 reveals why these enzymes are so big—many of the larger ones contain multiple types of catalytic modules as well as other functional modules as an integral part of a single polypeptide chain [see Table I in (Bayer et al. 1998c) 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 cellulosedegrading 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

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Fig. 6.20 Clostridium cellulolyticum enzyme system. An example of a cellulosomal system. Key to symbols: GH glycoside hydrolase (e.g., cellulase, xylanase, mannanase), CBM carbohydrate-binding module, Doc dockerin module

include Cel48S, and various Family-9 cellulases. The Cel48S subunit is a member of the Family-48 glycoside hydrolases, and this particular family is recognized as a critical component of bacterial cellulosomes (Morag et al. 1991, 1993; Wang et al. 1993, 1994; Wu et al. 1988). Several other processive cellulases are members of the Family-9 glycoside hydrolases. Cel9F and Cel9N are Theme-B Family-9 enzymes (> Fig. 6.7; Navarro et al. 1991). The other two are remarkably similar Theme-D enzymes, which exhibit nearly 95 % similarity along their common regions (Kataeva et al. 1999a, b; Zverlov et al. 1998c, 1999). The main difference between Cbh9A and Cel9K is the presence in the former of three extra modules (a Family-3 CBM and two modules of unknown function) (Kataeva et al. 2002, 2003, 2004, 2005). 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

6

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, numerous classic xylanases, i.e., those belonging to glycoside hydrolase families 10 and 11. In addition, two of the larger enzymes, Cel26H and Cel9/44J, contain hemicellulase components, i.e., Family-26 and Family-44 catalytic modules (a mannanase and a xylanase, respectively), together with a standard Family-5 and Family-9 (respectively) cellulase module in the same polypeptide chain (Ahsan et al. 1996; Yagu¨e et al. 1990). It is also interesting to note the presence of carbohydrate esterases together with xylanase modules in some of the enzyme subunits (i.e., XynU/A, XynY, XynZ and Cel5E), 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 Family-18 chitinase. The non-cellulosomal enzymes include another Theme-B Family-9 cellulase (Cel9I), and cell-bound forms of a xylanase (Xyn10X) and a lichenase (Lic16A), both of which contain multiple CBMs adjacent to the catalytic module. An additional non-cellulosomal Family-48 cellulase, Cel48Y, has also been described (Berger et al. 2007; Vazana et al. 2010). In the midst of all this complexity, the C. thermocellum non-cellulosomal cellulase system includes a simple Family-5 cellulase, Cel5C, which is completely devoid of additional accessory modules (Zverlov et al. 2005a; Feinberg et al. 2011). 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 (Lic16B) and chitinase (Chi18A) are also intriguing components of the cellulosome (Zverlov et al. 1991, 1998, 2002, 2005). 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. Subsequent genome sequencing of various strains of C. thermocellum has enhanced our understanding of the full complement of cellulosomal and non-cellulosomal enzymes produced by this bacterium (Zverlov et al. 2005a; Feinberg et al. 2011). Dockerin-containing components that are not directly involved in degradation of plant cell wall polysaccharides have

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Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

. Fig. 6.21 Clostridium cellulovorans: A second cellulosomal system. Key to symbols: GH glycoside hydrolase (e.g., cellulase, xylanase, mannanase), CBM carbohydrate-binding module, Doc dockerin module, SLH S-layer homology (module), Ig immunoglobulin-like domain, X domain of unknown function

also been identified (Kang et al. 2006; Schwarz and Zverlov 2006; Meguro et al. 2011). It is clear that the components and functions of the cellulosome system are much more complex than originally considered.

Gene Regulation of Cellulosomal Components Over the past 10 years, the genomic revolution has provided the complete sequence of numerous bacterial genomes. Recent analysis of 1,500 of these genomes indicated that 40 % of the genomes encode for at least one cellulase gene (Medie et al. 2012). Within the cellulosome-producing bacteria, there are dozens of different cellulosome-related genes, and their expression appears to be highly regulated. Our ability to elucidate the regulatory mechanisms have changed dramatically in recent years due to the availability of new genomic sequences, the development of genetic tools for some of the classical cellulosome-producing strains and the establishment of workable proteomic procedures which allow the identification and quantification of numerous gene products in

a single experiment. Much of the incentive for elucidating the regulatory mechanisms of cellulosome-producing bacteria is connected to their industrial potential for solubilizing lignocellulose for bioenergy production. In the context of this chapter, we will concentrate on new findings in C. thermocellum.

Regulation of Cellulase and Cellulosomal Genes in C. thermocellum The various cellulosomal genes in C. thermocellum are, for the most part, mono-cistronic and scattered throughout the chromosome (Brown et al. 2007; Raman et al. 2009). Since the number of known dockerin-bearing enzymes is almost ten times the number of cohesins in the scaffoldin subunit, a unique interaction between cohesin-dockerin pairs is unlikely. This has indeed been substantiated for C. thermocellum in which all of the scaffoldin-borne cohesins recognize nearly all of the dockerin-containing enzymes. Thus, the composition of the cellulosome is governed by the relative amounts of

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

6

. Fig. 6.22 Clostridium thermocellum: A complex cellulosomal system. Key to symbols: GH glycoside hydrolase (e.g., cellulase, xylanase, mannanase), CE carbohydrate esterase (e.g., acetyl xylan esterase and ferulic acid esterase), CBM carbohydrate-binding module, Doc dockerin module, SLH S-layer homology (module), Ig immunoglobulin-like domain, X domain of unknown function

the available dockerin-containing polypeptides that can be incorporated randomly into the complex (Bassen et al. 1995; Mitchell 1998). Regulation studies in C. thermocellum have indicated that the level and composition of the cellulosomal proteins vary with the composition of the growth media and cellobiose availability (Johnson et al. 1982; Nochur et al. 1990; Mishara et al. 1991; Bhat and Wood 1992; Nochur et al. 1992a, b; 1993; Raman et al. 2009). Recent studies (Dror et al. 2003a, b; 2005; Stevenson and Weimer 2005; Zhang and Lynd 2005; Brown et al. 2007; Raman et al. 2011; Riederer et al. 2011) have demonstrated that expression of many cellulose-related genes is influenced by growth rate and the presence of extracellular polysaccharides. The molecular regulatory mechanisms in C. thermocellum were, until recently, very much obscure, and the bacterium does not appear to encode many of the well-characterized global regulatory elements found in Gram-positive bacteria, including the pleiotropic regulator CodY (Sonenshein 2007). In this regard, one of the LacI homologues, GlyR3, was shown to be a negative regulator of celC, a non-cellulosomal cellulase gene, and laminaribiose (a b-1-3 linked glucose dimer) appears to be its molecular

inducer (Newcomb and Wu 2004; Demain et al. 2005; Newcomb et al. 2011). Remarkably, this was the first cellulose-related transcriptional factor identified in C. thermocellum.

Regulating by Sensing and the Involvement of Alternative Sigma Factors As outlined above, it was postulated that C. thermocellum must possess a regulatory system that allows it to sense and react to the presence of high-molecular-weight polysaccharides in the extracellular environment presumably without importing their lowmolecular-weight degradation products. While searching the C. thermocellum genome for the presence of carbohydrate binding modules, several Family-3 CBMs (CBM3s) were observed that were part of undefined polypeptides annotated as hypothetical proteins or membrane-associated proteins. Bioinformatic examination of these hypothetical peptides indicated possible homology to membrane-associated anti-s factors. Following this initial observation, searching the public nucleotide and protein databases revealed that three strains of

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Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

C. thermocellum contain a unique set of multiple ORFs resembling both Bacillus subtilis sigI and rsgI genes known to encode an alternative sI factor and its negative membrane-associated regulator RsgI, respectively (Asai et al. 2007). Bioinformatic analysis of over 1,200 bacterial genomes revealed that the C. thermocellum RsgI-like proteins are unique to this species and are not present in several other cellulolytic clostridial species (e.g., Clostridium cellulolyticum and Clostridium papyrosolvens) (Kahel-Raifer et al. 2010). However, several new genome sequences of other cellulosome-producing bacteria, e.g., Acetivibrio cellulolyticus and Clostridium clariflavum, have revealed similar types of multiple biomass-sensing systems. Indeed, the possible involvement of alternative s-factors in C. thermocellum was already suggested over 20 years ago by Mishara et al., following transcript analysis of three cellulosomal genes (celA, celD, celF) (Mishara et al. 1991). The C. thermocellum putative alternative s factors are homologous to the recently characterized sI gene in B. subtilis (Zuber et al. 2001; Asai et al. 2007; Schirner and Errington 2009). Each of the genes encoding sI-like factors is positioned adjacent to a downstream gene encoding a multi-modular protein that contains only one strongly predicted trans-membrane helix (TMD) (> Fig. 6.23). The 165-residue N-terminus of these trans-membrane proteins is homologous to the N-terminal

segment of the B. subtilis anti-sI factor, RsgI. The C-terminal modules of these RsgI-like proteins, purportedly located outside the cell membrane, contain predicted polysaccharide-related functions including carbohydrate-binding modules (CBM3, CBM42), sugar-binding elements (PA14), and glycoside hydrolase modules of families 10 and 5. The functional properties of the various elements of this system were verified experimentally (Kahel-Raifer et al. 2010; Nataf et al. 2010; Bahari et al. 2011). The binding properties of the extracellular sensing modules have been established with various polysaccharides including pectin, cellulose, arabinoxylan, and xylan (Kahel-Raifer et al. 2010; Bahari et al. 2011). Using isothermal titration calorimetry (ITC), it was possible to determine binding specificity and the dissociation constants (in the range of 0.02–1 mM) between the putative anti-sI factors to their corresponding s factors (Nataf et al. 2010). The expression of the relevant alternative s factor genes increased 3- to 30-fold in the presence of cellulose and xylan in the growth media, thus connecting their expression to direct detection of their extracellular polysaccharide substrates. Finally, the ability of sI1 to direct transcription from the sI1 promoter and from the promoter of celS (that encodes the Family 48 cellulase, Cel48S) was demonstrated in vitro by runoff transcription assays (Nataf et al. 2010).

. Fig. 6.23 Alternative s-factor operons in C. thermocellum. The operons are made of two genes, the s-factor gene (sig) and a transmembrane protein with an intracellular anti-s factor at the N-terminus, followed by a transmembrane domain (TMD) and an extracellular sensor module at the C-terminus. Many of these proteins contain carbohydrate-related modules: i.e., a CBM, a glycoside hydrolase (GH), or sugar-binding proteins (PA)

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

Since many alternative sigma factors auto-regulate their own expression, it is possible to identify the signature of their promoter sequence by determining the transcriptional start sites of the target genes. Using this approach, over 60 cellulosomal genes were assigned to their corresponding alternative sigma factor. In view of the above observations, a plausible model was proposed whereby the extracellular CBMs of putative anti-sIlike proteins can serve as biosensors that help assess the status of the biomass in the extracellular medium (> Fig. 6.24). When the target substrate is unavailable, the sI-like factor is attached to the N-terminal cytoplasmic domain of the RsgI-like protein. Upon interaction with the target polysaccharide, the corresponding RsgI-borne CBM may undergo a conformational change, leading to the release of the sI factor, which then associates with RNA polymerase. Target gene(s) are then transcribed including those that code for various carbohydrateactive enzymes (CAZymes) and cellulosomal scaffoldins, as well as the sI/RsgI-like operon itself. These various CBMs recognize and bind different plant cell wall polysaccharides,

6

which then induce different sets of CAZyme genes, thus activating the synthesis of the relevant glycoside hydrolases, carbohydrate esterases, and/or polysaccharide lyases.

New Genetic Tools for C. thermocellum One of the major obstacles in studying gene regulation in C. thermocellum was the lack of reliable transformation procedures and genetic tools. Recently, the laboratory of Prof. Lee Lynd from Dartmouth College developed two procedures for obtaining knockout mutants in C. thermocellum (Olson et al. 2010; Tripathi et al. 2010; Argyros et al. 2011). Both procedures allow selection and counter selection for an integration event. The first system uses the elegant approach devised initially for yeast, taking advantage of the fact that mutants lacking orotidine-50 -phosphate decarboxylase (Pyr) not only require uracil for their growth (uracil auxotrophs) but are also resistant to the pyrimidine analog 5-fluoro-orotic acid (5-FOA) (Boeke et al. 1984). Thus, when working with a background-strain lacking orotidine-50 -phosphate decarboxylase (the pyrF gene in C. thermocellum), the presence of the pyrF gene can be selected for or against simply by the inclusion of uracil or 5-FOA, respectively, in the growth medium (Kondo et al. 1991; Schneider et al. 2005). The second approach utilizes the activity of hypoxanthine phosphoribosyl transferase (Hpt), which is required for purine metabolism and makes purine antimetabolites, such as 8-azahypoxanthine (AZH), toxic. Another component of the system is the gene thymidine kinase (Tdk) (which conveniently C. thermocellum lacks). Tdk converts fluoro deoxyuracil (FUDR) to fluoro-dUMP which is a suicide inhibitor of thymidylate synthetase, and this can be used for counter selection in the presence of FUDR. This second approach allows obtaining multiple deletion mutants without the presence of selection markers.

Genomics and Metagenomics

. Fig. 6.24 Proposed mechanism for the activation of s factors by extracellular polysaccharides. The carbohydrate-binding sensing module – the CBM – is positioned on the outer surface of the bacterium and linked via a short transmembrane domain to a short anti-s peptide that binds and inactivates its cognate s-factor (off state). In the presence of various target polysaccharides, the CBM binds the polymers which induces a conformational change that results in the release of the s-factor, which now can initiate transcription of cellulose-utilization related promoters

In the past decade, major strides for enzyme discovery have been achieved by genomic and metagenomic approaches, combined with bioinformatic analyses. An early work on a bacterial genome involved a plant cell wall polysaccharide-degrading species (Nelson et al. 1999). This initial work was eventually followed by genome sequencing studies of additional cellulolytic bacteria (Lykidis et al. 2007; Xie et al. 2007; Berg Miller et al. 2009; Kataeva et al. 2009; Hemme et al. 2010; Morrison et al. 2010; Tamaru et al. 2010; Feinberg et al. 2011). Combined bioinformatics, proteomics, and transcriptomics characterization can serve to reveal the components of the relevant enzyme system(s) in a given bacterium (Marcotte et al. 1999; Zverlov et al. 2005; Flint et al. 2008; Li et al. 2009; Raman et al. 2009; 2011; Rincon et al. 2010; Brulc et al. 2011; Dam et al. 2011). The metagenomic approach utilizes genetic material directly from complex natural ecosystems, rather than using cultivated

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cells (Schloss and Handelsman 2003; Handelsman 2004). The advantage of this approach is that bias against unculturable bacteria is avoided and enzyme discovery is more representative. However, metagenomic libraries can introduce new types of biases due to nonuniform recovery of inserts and large numbers of clones required to cover the metagenome. Metagenomic analyses of different cellulosecontaining ecosystems can serve to provide insight into novel types of enzymes that can be used for degradation of plant cell wall polysaccharides (Cottrell et al. 2005; Ferrer et al. 2005; Warnecke et al. 2007; Brulc et al. 2009; Li et al. 2009; Berg Miller et al. 2012). > Table 6.1 provides a list of cellulose-degrading bacteria whose genomes have been sequenced. It is clear that new genome sequences of cellulase- and cellulosome-producing bacteria will continue to accumulate, at least until the current involvement with cellulosic biomass-to-biofuel efforts remains in vogue.

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 cellulosomes 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 a legacy, inherently encoded in the sequences of the functional modules 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 glycoside hydrolases.

Horizontal Gene Transfer It is clear that very similar enzymes which comprise a given glycoside 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 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., CBMs, 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 module 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 glycoside 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 modules 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 glycoside hydrolases for a given family of crystalline cellulose-binding CBM, which is entirely independent of the type of catalytic module borne by the complete enzyme. In this 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 CBM, irrespective of the type of catalytic module. In contrast, those of other bacteria, e.g., Bacillus subtilis, Caldocellum saccharolyticum, Erwinia carotovora, and various clostridia, appear to prefer Family-3 CBMs. Moreover, the position of the CBM in the gene may be different for different genes. For example, the CBM may occur upstream or downstream from the catalytic module; 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 modules can be transferred en

Lignocellulose-Decomposing Bacteria and Their Enzyme Systems

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 duplication which would propagate the insertion of repeated modules, e.g., the multiple cohesin modules in the scaffoldins, or even smaller units, such as the linker sequences or the duplicated calciumbinding loop of the dockerin module. Domain shuffling can account for the observed permutations in the arrangement of modules in scaffoldin subunits from different species (> Fig. 6.15). 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 noncellulosomal enzyme systems, whereas phylogenetically related organisms that inhabit similar niches may be characterized by different types of enzyme systems. It is clear that in order 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—in order to sharpen existing notions regarding the enzymatic degradation of plant cell wall polysaccharides or to formulate new ones.

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Acknowledgments Grants from the Israel Science Foundation (administered by the Israel Academy of Sciences and Humanities, Jerusalem), the US-Israel Binational Foundation (BSF), the Israel Ministry of Science (IMOS), and by the Weizmann Institute of Science Alternative Energy Research Initiative (AERI) are greatly appreciated. The authors are also pleased to acknowledge the establishment of an Israeli Center of Research Excellence (I-CORE) managed by the Israel Science Foundation (grant No 152/11) and additional support by the Technion-Niedersachsen Research Cooperation Program. Y.S. holds the Erwin and Rosl Pollak Chair in Biotechnology at the Technion, E.A.B. is the incumbent of The Maynard I., and Elaine Wishner Chair of Bio-organic Chemistry.

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7 Aerobic Methylotrophic Prokaryotes Ludmila Chistoserdova1 . Mary E. Lidstrom2 1 Department of Chemical Engineering, University of Washington, Seattle, WA, USA 2 Department of Chemical Engineering and Department of Microbiology, University of Washington, Seattle, WA, USA

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 Dissimilatory Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 Methane Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 269 Methanol Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Quinoprotein Methanol Dehydrogenases . . . . . . . . . . . . . . . 270 NAD-Linked Methanol Dehydrogenases . . . . . . . . . . . . . . . . 270 Methanol: N,N0 -dimethyl-4-nitrosoaniline Oxidoreductase (MNO) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Oxidation of Methylated Amines . . . . . . . . . . . . . . . . . . . . . . . 271 Trimethylamine and Dimethylamine . . . . . . . . . . . . . . . . . . . . 271 Methylamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 271 Utilization of Methylated Sulfur Species . . . . . . . . . . . . . . . . 272 Utilization of Halomethanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 Formaldehyde Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 The Tetrahydromethanopterin-Linked Formaldehyde Oxidation Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 The Tetrahydrofolate-Linked C1 Transfer Pathway and Its Dual Role in Methylotrophy . . . . . . . . . . . . . . . . . . . . 273 NAD- and Mycothiol-Linked Formaldehyde Dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274 NAD- and GSH-Linked Formaldehyde Dehydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 274 NAD-Linked Formaldehyde Dehydrogenase . . . . . . . . . . . . 274 Cyclic Formaldehyde Oxidation Pathway . . . . . . . . . . . . . . . 275 Distribution of Formaldehyde Oxidation Pathways in Methylotrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Formate Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Assimilatory Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276 Serine Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276 Glyoxylate Regeneration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 277 Ribulose Monophosphate Cycle . . . . . . . . . . . . . . . . . . . . . . . . . 278 The CBB Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 278 Methylotrophic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 Methanotrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 Non-methane-Utilizing Methylotrophs . . . . . . . . . . . . . . . . . 279 Genetics in Aerobic Methylotrophs . . . . . . . . . . . . . . . . . . . . . . . . 279 Genetic Capabilities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 Methylotroph Genomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 280 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 280

Abstract This chapter describes biochemical pathways operating in aerobic methylotrophic bacteria. We first define aerobic methylotrophy as a specific metabolic capability and describe the phylogenetic diversity of these bacteria. We then describe enzymes involved in primary oxidation of different single carbon substrates, resulting in formaldehyde or methyl or methylene radical, and describe the variety of pathways used for their assimilation and dissimilation. We also give a brief account of genetic manipulation tools in methylotrophic bacteria and examples of systems approaches for studying their metabolism, including availability of whole genome sequence information.

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 widespread in nature, being found in a variety of aquatic and terrestrial habitats (King 1992; Trotsenko and Murrell 2008), including extreme environments (Pol et al. 2007; Dunfield et al. 2007; Islam et al. 2008; Antony et al. 2010). 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 microbes are known, such as methanotrophic archaea (Knittel and Boetius 2009), methanoland methylamine-utilizing methanogenic archaea (Thauer 1998), and methylotrophic clostridia (Ragsdale and Pierce 2008), this chapter will cover only the aerobic methylotrophs. However, an exception will be made for the newly described methylotrophs of the NC10 phylum that favor an anaerobic lifestyle but require oxygen for methane oxidation and utilize metabolic pathways common with aerobic methylotrophs (Ettwig et al. 2010). > Table 7.1 lists the major known groups of aerobic methylotrophs with examples of family affiliations and genome sequence information.

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_68, # Springer-Verlag Berlin Heidelberg 2013

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. Table 7.1 Major groups of methylotrophs showing family affiliations, major assimilatory pathways, and genome sequence availability. Secondary assimilatory pathways are noted in parentheses Phylum/class

Family

Major assimilation pathway Representative genome references

Methane utilizers (methanotrophs) Gammaproteobacteria

Methylococcaceae

RuMP (serine, CBB)

Ward et al. (2004); Boden et al. (2011); Svenning et al. (2011); Vuilleumier et al. (2012)

Alphaproteobacteria

Methylocystaceae

Serine

Stein et al. (2010), (2011)

Beijerinckiacaee

Serine (CBB)

Chen et al. (2010a)

Verrucomicrobia

Methylacidiphilaceae CBB

Candidate phylum NC10 Not formalized

CBB

Hou et al. (2008) Ettwig et al. (2010)

Non-methane utilizers Alphaproteobacteria

Betaproteobacteria

Methylobacteriaceae Serine

Vuilleumier et al. (2009); Marx et al., (2012) for Methylobacteriaceae

Hyphomicrobiaceae

Serine

Vuilleumier et al. (2011); Brown et al. (2011)

Bradyrhizobiaceae

Serine

None

Acetobacteraceae

Serine

Greenberg et al. (2007)

Rhodobacteraceae

CBB

Li et al. (2011); Siddavattam et al. (2011)

Xanthobacteraceae

CBB

JGI

Methylophilaceae

RuMP

Chistoserdova et al. (2007a); Giovannoni et al. (2008); Lapidus et al. (2011)

Burkholderiaceae

Serine (CBB)

JGI

Rhodocyclaceae

Serine (CBB)

Kittichotirat et al. (2011)

Unclassified

Serine (CBB)

Kane et al. (2007)

Gammaproteobacteria

Piscirickettsiaceae

RuMP

Boden et al. (2011); Han et al. (2011)

Gram-positive

Bacillaceae

RuMP

Heggeset et al. (2012)

Pseudonocardiaceae RuMP

None

Micrococcaceae

JGI

RuMP

RuMP, serine, or CBB, utilize, respectively, ribulose monophosphate, serine or Calvin–Benson–Bassham cycles for C1 assimilation. JGI, genome information is available prior to formal publication via the Joint Genome Institute web sites: http://img.jgi.doe.gov/cgi-bin/w/main.cgi; http://genome.jgi-psf.org/

Aerobic methylotrophic bacteria are phylogenetically diverse, with representatives found among the Alpha-, Beta-, and Gammaproteobacteria and Verrucomicrobia as well as the high and low G + C Gram-positive bacteria (Firmicutes; > Table 7.1). Methanotrophs of the NC10 phylum carry out methylotrophy in a false anaerobic fashion, by producing oxygen intracellularly (intra-aerobic metabolism; Ettwig et al. 2010). Aerobic methylotrophs were formerly divided into obligately methylotrophic species, that is, species incapable of growing on any compounds containing carbon–carbon bonds and facultatively methylotrophic species, that is, species that can grow on a limited or a large number of multicarbon compounds (Anthony 1982). Recently, both the discovery of novel species of methylotrophs and the cultivation techniques more closely reflecting environmental conditions, including adjusted expectations for growth rates, have questioned whether this separation is valid, as facultative methylotrophs have been identified in groups previously known to contain only obligate methylotrophs (Dedysh et al. 2005; Dunfield et al. 2010; Im and Semrau 2011; Kalyuzhnaya et al. 2011). A special functional group is distinguished within methylotrophs, called

methanotrophs, the organisms capable of growth on methane, and these are important participants in the global methane cycle (Anthony 1982; King 1992; Trotsenko and Murrell 2008). Some of the methylotrophs also can use N2 as a nitrogen source and therefore are considered to be diazotrophs (Anthony 1982; Trotsenko and Murrell 2008). 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. A number of variations of these metabolic pathways have been identified. > Figure 7.1 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

Aerobic Methylotrophic Prokaryotes

1 CH3SO3H

3 (CH3)2NH

8 CH3OH

DMSP 10

(CH3)3N

CH2Cl2 CH2Br2

CH4

9

4

DMS 11

CH3Cl CH3Br

CH3NH2

2

6

5

CH2O 13

14

CH2=H4F

7

DMSP 12

15 RuMP cycle

HCOOH 16

Biomass

CO2

Serine cycle Biomass

CBB cycle Biomass

. Fig. 7.1 Metabolism of one-carbon compounds in aerobic methylotrophic bacteria. 1 methane monooxygenase, 2 methanol dehydrogenase, 3 trimethylamine dehydrogenase or trimethylamine monooxygenase, 4 dimethylamine dehydrogenase or dimethylamine monooxygenase, 5 methylamine dehydrogenase, 6 N-methylglutamate pathway, 7 halomethane methyltransferase, 8 dihalomethane dehalogenase, 9 methanesulfonate monooxygenase, 10 dimethylsulfoniopropionate (DMSP) lyase, 11 dimethylsulfide monooxygenase, 12 DMSP demethylase, 13 formaldehyde oxidation systems, 14 methylene H4F oxidation pathway, 15 assimilatory H4F-linked C1 transfer pathway, and 16 formate dehydrogenase enzymes (Adapted from Chistoserdova (2011))

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. Some C1 substrates are degraded via demethylation, resulting in transfer of methyl groups to tetrahydrofolate, thus not producing formaldehyde (> Fig. 7.1). Other methylotrophs are capable of growth on reduced C1 compounds by oxidizing them to CO2 and then assimilating the CO2 via the classical Calvin–Benson–Bassham cycle. These are known as autotrophic methylotrophs (Anthony 1982). The diagram shown in > Fig. 7.1 is an amalgam of the known diversity of methylotrophic metabolism, and no single known methylotroph can carry out all of these types of metabolism. However, the concept of ‘‘modularity’’ of methylotrophy suggests that, while types of methylotrophs exist possessing only a minimal set of methylotrophy metabolic modules (Giovannoni et al. 2008), methylotrophs may also exist that are omnipotent in their metabolic capabilities (Chistoserdova 2011).

Dissimilatory Metabolism Aerobic methylotrophs contain specialized pathways for dissimilatory metabolism during methylotrophic growth. In general,

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the primary methylotrophic substrates are oxidized, demethylated, or dehalogenated to the level of formaldehyde, methyl-, or methylene radical (typically transferred onto tetrahydrofolate; H4F) by specialized oxidases, dehydrogenases, methyltransferases, or dehalogenases. The dehydrogenases are generally coupled to energy metabolism at the level of cytochromes, while the oxidases, methyltransferases, and dehalogenases are usually nonenergy conserving. Formaldehyde (methyl-, methylene-H4F) is then further oxidized to the formyl level by one of a number of the oxidation systems, which usually generate a reduced pyridine nucleotide. Carbon at the level of formate is then oxidized to CO2 via formate dehydrogenases, some of which also generate reduced pyridine nucleotides (Laukel et al. 2003; Chistoserdova et al. 2004b), while others may link directly into the electron transport chain (Chistoserdova et al. 2007a).

Methane Oxidation The enzyme that oxidizes methane to methanol in the methanotrophic bacteria is a mixed-function oxidase called ‘‘methane monooxygenase’’ (MMO), which incorporates one atom of oxygen from O2 into methane and requires reducing power to reduce the second oxygen atom of O2 to H2O. 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; Trotsenko and Murrell 2008). The pMMO appears to be the enzyme almost universally widespread among methanotrophs, while the sMMO is only found in some organisms, typically in addition to pMMO, a single exception so far being members of the family Beijerinckiaceae that only possess sMMO (Dedysh et al. 2000; Chen et al. 2010a; Vorobev et al. 2011). The sMMO has been purified from both alpha- and gammaproteobacterial methanotrophs (Lipscomb 1994), and it is similar in all cases. It consists of three components: a hydroxylase (consisting of three polypeptides and a nonheme iron center), component B (with no cofactors), and a reductase that contains FAD and a Fe2S2 cluster (Lipscomb 1994). The sMMO uses NADH as a source of reducing power and contains a hydroxo-bridged diiron 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 regarding 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 (Rosenzweig et al. 1993, 1997; Elango et al. 1997; Chang et al. 1999; Walters et al. 1999). Genes encoding the subunits of the sMMO (mmo genes) are typically organized into clusters on the chromosomes, and high similarity at the amino acid level is typically found

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between different species (Ward et al. 2004; Chen et al. 2010a; Stein et al. 2010, 2011). The pMMO has also been purified in an active state from both alpha- and gammaproteobacterial methanotrophs (Zahn and DiSpirito 1996; Nguyen et al. 1998). The pMMO has a narrower substrate range than the sMMO (Hanson and Hanson 1996; Hakemian and Rosenzweig 2007). The enzyme is encoded by a conserved gene cluster pmoCAB. Some genomes contain a single pmoCAB cluster (Stein et al. 2011), while in others, multiple, sometimes divergent, copies are present (Ward et al. 2004; Hou et al. 2008). Crystal structures of pMMO enzymes from different bacteria have also been determined (Lieberman and Rosenzweig 2005; Hakemian et al. 2008), and the role of a dicopper center in the catalysis has been unequivocally proven (Balasubramanian et al. 2010). The importance of copper in pMMO-mediated methane oxidation is also highlighted by the recent discovery of methanobactins, highaffinity copper-binding molecules, akin to siderophores that appear to be involved in copper uptake, regulation of methane monooxygenase expression, protection against copper toxicity, and particulate methane monooxygenase activity (Kim et al. 2004; Balasubramanian and Rosenzweig 2008). The source of reducing power for this enzyme in vivo is still not defined. 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, sMMO is expressed. It has been demonstrated that this regulation occurs primarily at the transcriptional level, but additional regulatory mechanisms are likely involved (Nielsen et al. 1997; Murrell et al. 2000; Hakemian and Rosenzweig 2007).

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 known classes of enzymes, quinoprotein methanol dehydrogenases (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,N0 -dimethyl-4-nitrosoaniline oxidoreductase (MNO) found in other Gram-positive strains (Bystrykh et al. 1993, 1997). In general, methanol oxidation is an energyconserving step, either generating reduced cytochromes or reduced pyridine nucleotides.

Quinoprotein Methanol Dehydrogenases Many of the known Gram-negative methanol- and methaneutilizing bacteria contain a periplasmic methanol dehydrogenase consisting of two different subunits forming an a2b2 structure, containing the cofactor pyrroloquinoline quinone (PQQ) and encoded by genes named mxaFI (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 possessing this enzyme 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). Recently, it has been demonstrated that some methylotrophs, specifically the methylotrophs belonging to the orders of Burkholderiales and Rhodocyclales, encode a different type of MDH, named MDH2 (Kalyuzhnaya et al. 2008a). This enzyme appears to be composed of a single type of subunit whose protein sequence shows very low similarity to MxaF proteins (less than 35 % amino acid identity). Instead, Mdh2 is highly similar (up to 80 % amino acid identity) to a class of PQQlinked dehydrogenases that includes alcohol dehydrogenases typically exhibiting low affinity for methanol (Kalyuzhnaya et al. 2008a). Mdh2 enzymes were proposed to have resulted from convergent evolution toward methanol oxidation capability, as opposed to diverging from MxaFI type of MDH enzymes. In addition, a number of methylotrophs have now been characterized that are capable of methanol metabolism but contain neither mxaFI nor mdh2 (Giovannoni et al. 2008; Hou et al. 2008; Kalyuzhnaya et al. 2011). These must possess a different type of MDH. One gene, named xoxF, encoding a polypeptide whose sequence is similar to the sequence of MxaF (approximately 50 % amino acid identity) has been proposed to encode an enzyme replacing the classic MDH (Wilson et al. 2008; Giovannoni et al. 2008; Hou et al. 2008). However, while a low MDH activity was demonstrated for purified XoxF (Schmidt et al. 2010), its role in methylotrophy has not been unequivocally proven.

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 this MDH shows identity with type II alcohol dehydrogenases (de Vries et al. 1992).

Methanol: N,N0 -dimethyl-4-nitrosoaniline Oxidoreductase (MNO) Other Gram-positive methylotrophs (Amycolatopsis and Mycobacterium) oxidize methanol via a methanol: N,N0 -dimethyl-4nitrosoaniline oxidoreductase (MNO), which is a decameric

Aerobic Methylotrophic Prokaryotes

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). The structural gene for this MDH shows identity with type III alcohol dehydrogenases (Nagy et al. 1995b; Park et al. 2010).

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 standard methylotrophic assimilatory and dissimilatory pathways (> Fig. 7.1).

Trimethylamine and Dimethylamine Trimethylamine is oxidized to dimethylamine and formaldehyde by trimethylamine dehydrogenase (> Fig. 7.1). This enzyme is a flavoprotein that also contains two Fe2S2 clusters and two molecules of ADP (McIntire 1990). Gene sequences suggest that trimethylamine and dimethylamine dehydrogenases are evolutionarily related (Yang et al. 1995). A second pathway for utilization of trimethylamine occurs in which a trimethylamine monooxygenase (TMM) oxidizes trimethylamine to trimethylamine N-oxide. The N-oxide is subsequently demethylated by trimethylamine demethylase to dimethylamine and formaldehyde (Anthony 1982). TMM is a flavoprotein and is similar in its properties to the well-characterized eukaryotic flavine-containing monooxygenases (Chen et al. 2011). Genes encoding TMM have been identified in a variety of bacteria, suggesting an important environmental role (Chen et al. 2011). Dimethylamine is oxidized to methylamine and formaldehyde by dimethylamine monooxygenase (> Fig. 7.1). While TMM can oxidize dimethylamine in vitro, it is proposed that this function must be carried out in vivo by a different enzyme (Chen et al. 2011).

Methylamine Three possible routes are known in bacteria for utilizing methylamine. The first of these involves the periplasmic enzyme, methylamine dehydrogenase (MADH), that is widespread among methylotrophic Proteobacteria (Anthony 1982). MADH is a quinoprotein shown to contain the cofactor tryptophan tryptophylquinone (TTQ). TTQ is formed by covalent

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cross-linking 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 (Pearson et al. 2003; Davidson 2005; Wilmot and Davidson 2009). The MADH converts primary amines to their corresponding aldehydes plus ammonia, and electrons are transferred to an electron acceptor. So far, three types of MADH-specific acceptors have been characterized or proposed: a small copper protein, amicyanin (Ferrari et al. 2004; Meschi et al. 2010), a copper protein azurin (Gak et al. 1995), or a small cytochrome (Kalyuzhnaya et al. 2008b). The electrons are further transferred to the respiratory chain via a c-type cytochrome (Ferrari et al. 2004; Meschi et al. 2010). Structural, kinetic, and site-directed mutagenesis studies have characterized protein-protein interactions and mechanisms of catalysis and electron transfer by TTQ in great detail (Wilmot and Davidson 2009; Jensen et al. 2010). The genes encoding the functions required for active MADH (mau genes) are coserved among alpha-, beta-, and gammaproteobacterial methylotrophs, found in a cluster mauBEDAGLMN, excepting the variations in the electron acceptor mentioned above. In some organisms, the mau gene cluster was found to be flanked by IS (insertion sequence) elements, suggesting that the methylamine oxidation capability may be a subject of lateral gene transfers (Vuilleumier et al. 2009). Indeed, there are a few examples of closely related methylotroph species differing with regard to presence of the mau genes (Kalyuzhnaya et al. 2008b; Vuilleumier et al. 2009), supporting this hypothesis. An alternative pathway used by Gram-negative methylotrophs is the indirect pathway involving the conversion of methylamine to N-methylglutamate, via N-methylglutamate synthase, and finally to methylene-H4F, via N-methylglutamate dehydrogenase (Latypova et al. 2010). Gene clusters encoding these two enzymes typically also encode g-glutamylmethylamide synthase, whose role remains poorly understood. In Methyloversatilis universalis, it was found nonessential and proposed to serve in balancing carbon/nitrogen flow (Latypova 2010), while in Methylocella silvestris, et al. g-glutamylmethylamide was found to be an essential intermediate (Chen et al. 2010c). In some methylotrophs, the N-methylglutamate pathway is the only pathway for methylamine oxidation (Latypova et al. 2010; Chen et al. 2010c), while in others, it is present along with MADH, playing a secondary role (Hendrickson et al. 2010). The NMG pathway is also present in non-methylotrophs, which employ it for utilization of methylamine as a nitrogen source (Chen et al. 2010b). Like MADH, this pathway has also been proposed to be a potential subject of lateral gene transfers (Chen et al. 2010c). In Gram-positive methylotrophs, represented by 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). 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).

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Utilization of Methylated Sulfur Species Some of the most environmentally important methylated sulfur species are dimethylsulfoniopropionate (DMSP), dimethylsulfide (DMS), and methanesulfonic acid (MSA; Kelly and Murrell 1999; Yoch 2002). Some organisms have been described that are also capable of utilizing dimethylsulfone and dimethylsulfoxide (DeBont et al. 1981; Suylen and Kuenen 1986; Kanagawa and Kelly 1986; De Zwart et al. 1996; Borodina et al. 2000). These substrates are first converted to DMS, by respective reductases (Borodina et al. 2002). DMS is then converted to methanethiol and formaldehyde by DMS monooxygenase (DmoAB; Boden et al. 2010) (> Fig. 7.1). 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. 7.1). The methanesulfonate-utilizing species oxidize this substrate to sulfite and formaldehyde by NADH-dependent methanesulfonate monooxygenase (MsmABCD; Kelly and Murrell 1999; Baxter et al. 2002). Microbial degradation of DMSP is known to proceed via two alternative routes, demethylation, catalyzed by DMSP-dependent demethylase (DmdA) and cleavage, catalyzed by a few different types of DMSP lyases producing DMS (Reisch et al. 2011). So far, degradation of DMSP has not been described as a bona fide case of methylotrophy. However, genomes of some of the DMSPdegrading bacteria encode methylotrophy pathways, suggesting that they may co-metabolize methyl groups using these pathways (Chistoserdova 2011).

Utilization of Halomethanes A number of methylotrophic bacteria are known that are capable of aerobic growth on halomethanes such as chloromethane, bromomethane, and dimethylchloride (Leisinger et al. 1994; Leisinger and Braus-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 alphaproteobacterial methylotrophs within a clade of rhizobia (Schaefer and Oremland 1999; Coulter et al. 1999). Dichloromethane degradation involves a glutathione-linked dehalogenase that produces formaldehyde (Leisinger et al. 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 et al. 1999). In Methylobacterium strain CM4, the methyltransferase reaction is coupled to H4F derivatives to produce formate, followed by formate oxidation (Vannelli et al. 1999; > Fig. 7.1). Assimilation occurs via methylene-H4F and the serine cycle. In strain CC495, which is one of the chloromethane utilizers that classes near rhizobia, evidence is presented for a disulfide-coupled reaction

in which methanethiol is the product (Coulter et al. 1999). In that case, it has been proposed that methanethiol is oxidized to formaldehyde, and metabolism proceeds by standard 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 and Dalton 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 formaldehydelimited chemostat by first establishing cultures on choline, then adding low levels of formaldehyde, and finally eliminating the choline gradually (Levering et al. 1986). However, adaptation of some Methylobacterium strains to extremely high formaldehyde concentrations has also been reported (Chongcharoen et al. 2005). A number of formaldehyde oxidation systems are known in methylotrophs. The simplest of these is formaldehyde dehydrogenase (FaDH), which converts formaldehyde to formate. A number of NAD-linked and dye-linked (presumably PQQcontaining and cytochrome-linked) FaDHs have been identified from methylotrophs, but the low activity and general lack of inducibility of these enzymes in most cases have called their physiological role into question (Hirt et al. 1978; Stirling and Dalton 1978; Anthony 1982; Marison and Attwood 1982; Weaver and Lidstrom 1985; Van Ophem and Duine 1990; Chistoserdova et al. 1991; Attwood et al. 1992). 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). However, a PQQ-linked FaDH from Methylococcus capsulatus has been described that appears to be specifically expressed in cells expressing pMMO (versus sMMO), and this was proposed to be the major formaldehydeoxidizing enzyme in this organism when grown in the presence of copper (Zahn et al. 2001). The respective gene is annotated in the genome of M. capsulatus as a sulfide-quinone reductase (Ward et al. 2004), and gene homologs are found across Proteobacteria. Below we describe pathways/enzymes with a proven function in formaldehyde oxidation.

The Tetrahydromethanopterin-Linked Formaldehyde Oxidation Pathway One of the most persistent formaldehyde oxidation pathways in methylotrophs of various groups is the pathway employing tetrahydromethanopterin (H4MPT) as a cofactor and encoded by at least 20 genes (Chistoserdova et al. 2009; Chistoserdova 2011; > Fig. 7.2a). This pathway is analogous to the pathway operating in the methanogenesis pathway in methanogenic

Aerobic Methylotrophic Prokaryotes

Primary oxidation

Primary demethylation

CH2O

CH3-H4F

1 CH2=H4MPT 2 A

CO2

10

B

CH≡H4MPT 3

CH≡H4F 10

CHO-H4MPT 4 5 HCOOH

CHO-H4F

6 CHO-H4F C

9 CH2=H4F

11 HCOOH 5 CO2

7 CH≡H4F 8 CH2=H4F Serine cycle

. Fig. 7.2 Pterine-linked pathways for formaldehyde oxidation. (a) tetrahydromethanopterin (H4MPT)-linked pathway for formaldehyde oxidation; (b) tetrahydrofolate (H4F)-linked C1 transfer pathway involving FolD; (c) H4F-lnked C1 transfer pathway involving MtdA and Fch. The latter operates in the reductive direction. 1 formaldehyde-activating enzyme, 2 methylene- H4MPT dehydrogenase (MtdB), 3 methenyl-H4MPT cyclohydrolase, 4 formyltransferase hydrolase complex, 5, formate dehydrogenase, 6 formyl-H4F ligase (FtfL), 7 methenylH4F cyclohydrolase, 8 methylene-H4F/methylene-H4MPT dehydrogenase (MtdA), 9 methyl-H4F reductase, 10 bifunctional methylene-H4F dehydrogenase/methenyl-H4F cyclohydrolase (FolD), 11 formyl-H4F hydrolase (PurU) (Adapted from Chistoserdova et al. (2009))

Archaea and to a pathway operating for formaldehyde oxidation in sulfate-reducing Archaea, such as Archaeoglobus fulgidus (therefore, the respective enzymes were initially called archaeal-like enzymes; Chistoserdova et al. 1998, 2004a). The significance of this pathway was first discovered in Methylobacterium extorquens AM1 (Chistoserdova et al. 1998), when it was demonstrated to be an indispensable pathway for both formaldehyde oxidation, with generation of NAD(P)H, and for formaldehyde detoxification (Hagemeier et al. 2000; Marx et al. 2003a). Mutants of M. extorquens with lesions in the genes involved in this pathway were all methylotrophy-negative, and some demonstrated a remarkable threshold for methanol sensitivity (as low as 90 mM while wild-type organism would grow at over 200 mM methanol; Chistoserdova et al. 2005a). While this pathway has been identified in most methylotrophs (Chistoserdova 2011), it does not play a crucial role in all of them. For example, in betaproteobacterial methylotrophs, this

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pathway appears to serve an auxiliary function, while the cyclic ribulose monophosphate pathway (see below) is the major pathway in oxidizing and detoxifying formaldehyde, and mutants with lesions in this pathway can be easily generated (Chistoserdova et al. 2000; Kalyuzhnaya et al. 2005; Hendrickson et al. 2010). Most of the archaeal-like enzymes in methylotrophs show significant identity to the corresponding archaeal enzymes, suggesting a common evolution (Chistoserdova et al. 1998, 2004a). However, some of the enzymes in the pathway are bacteria-specific. Notably, the archaeal H2- or F420-enzymes that interconvert methylene- and methenyl-H4MPT are not found in the aerobic methylotrophs or other bacteria possessing this pathway. Instead, two enzymes have been identified that oxidize methylene-H4MPT to methenyl-H4MPT: MtdA and MtdB (Vorholt et al. 1998; Hagemeier et al. 2000). MtdA will only use NADP as a cofactor but will oxidize both methylene-H4MPT and methylene-H4F (Vorholt et al. 1998), while MtdB will use either NAD or NADP but is specific to H4MPT (Hagemeier et al. 2000). MtdA and MtdB are distant homologs (approximately 30 % amino acid identity; Chistoserdova et al. 1998; Vorholt et al. 1998; Hagemeier et al. 2000). The available data suggest so far that MtdB is the main enzyme in the H4MPT-linked formaldehyde oxidation pathway and MtdA cannot substitute for MtdB (Chistoserdova et al. 1998). In addition to MtdA and MtdB, a novel class of Mtd enzymes has been recently described. Some of these have been classified as MtdC, and these represent the division Planctomycetes and a novel deeply divergent phylum of uncultivated bacteria from Lake Washington, termed Phylum LW (Chistoserdova 2011). These enzymes have high affinities for methylene-H4MPT and NADP but low affinities for methylene H4F or NAD, distinguishing them from MtdA and MtdB enzymes (Vorholt et al. 2005). Accordingly, it has been demonstrated that MtdC could not functionally substitute for either MtdA or MtdB (Vorholt et al. 2005). Thus, while phylogenetically more related to MtdA, MtdC must fulfill a function more similar to the function of MtdB, as part of the H4MPT-linked pathway for formaldehyde oxidation/detoxification (Vorholt et al. 2005). However, it remains unknown whether MtdC is involved in methylotrophy. Analysis of the recently available genomic sequences suggests that the diversity of Mtd enzymes must extend even further. For example, genes predicted to encode divergent Mtd proteins have been identified in the genomes of Methylophaga thiooxidans, Nitrosococcus halophilus, and Anaerobaculum hydrogeniformans, and these may represent new classes and may express different sets of substrate specificities, based on their phylogenetic positions (Chistoserdova 2011).

The Tetrahydrofolate-Linked C1 Transfer Pathway and Its Dual Role in Methylotrophy A C1 transfer pathway linked to tetrahydrofolate (H4F), analogous to the H4MPT-linked pathway, exists in many

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methylotrophs (and across all life). In most organisms, including some methylotrophs, this pathway involves FolD, a bifunctional enzyme possessing methylene-H4F dehydrogenase and methenyl-H4F cyclohydrolase activities. In methylotrophs that grow on compounds whose primary degradation involves demethylation reactions, such as chloromethane-degrading Methylobacterium extorquens (formerly chloromethanicum) CM4, FolD appears to be specifically involved in dissimilation of these compounds, along with respective methyltransferases (CmuAB), methyl-H4F reductase (MetF), and formyl-H4F hydrolase (PurU; Vannelli et al. 1999; > Fig. 7.2b). However, some methylotrophs, for example, other M. extorquens strains and Methylococcus capsulatus, do not posses FolD. Instead, they employ two alternative enzymes in this pathway, MtdA and Fch, responsible for, respectively, methylene-H4F dehydrogenase and methenyl-H4F cyclohydrolase activities. These enzymes have been characterized in M. extorquens AM1, and, based on enzyme kinetics (Vorholt et al. 1998; Pomper et al. 1999), mutant phenotypes (Chistoserdova and Lidstrom 1994; Marx and lidstrom 2004) and metabolite flux analysis (Marx et al. 2005; Crowther et al. 2008), they appear to be involved in a pathway that acts in a reductive rather than oxidative direction in this organism, that is, this pathway enables transfer of C1 units into the serine cycle using H4F as an adduct (Crowther et al. 2008; > Fig. 7.2c). The latter study thus challenged the long-standing dogma of spontaneous reaction between formaldehyde and H4F being of physiological significance. The difference in physiological roles of MtdA and FolD is highlighted by mutant complementation experiments: While MtdA could act as a functional substitute for FolD (Studer et al. 2002), FolD could not complement a lesion in MtdA (Marx and Lidstrom 2004). So far, the significance of the H4MPT-linked activity of MtdA remains unclear as it cannot substitute for MtdB in the H4MPT-linked oxidative pathway (Chistoserdova et al. 1998), but it suggests that this enzyme may have an additional, not yet identified in vivo function. Understanding the specific functions of different variants of H4F-utilizing C1 transfer pathways is further complicated by the use of two alternative enzymes for transferring C1 units between the levels of formyl-H4F and formate: formyl-H4F ligase (FtfL) being a reversible enzyme (Marx et al. 2003b) and formyl-H4F hydrolase (PurU) being a nonreversible enzyme (Nagy et al. 1995a). Thus, in the organisms possessing FtfL, the pathway can in principle operate in both directions, while in the organisms possessing only PurU, the pathway can only operate in the oxidative direction. In keeping with these characteristics, the organisms utilizing the MtdA/Fch variant of the pathway always employ FtfL, while organisms utilizing the FolD variant can employ either FtfL or PurU or both (Chistoserdova 2011). A few methylotrophs, exemplified by Methylibium petroleiphilum and Methylobacterium extorquens CM4, encode both variants of the pathway (Vannelli et al. 1999; Studer et al. 2002; Kane et al. 2007). In the latter, PurU was demonstrated to specifically operate in degradation of chloromethane, and FtfL cannot functionally substitute for PurU

(Vannelli et al. 1999; Studer et al. 2002). However, from the experimental data available, it is not clear whether FtfL is simply not present at sufficient levels under these growth conditions or whether it can only operate in the reductive direction in this organism.

NAD- and Mycothiol-Linked Formaldehyde Dehydrogenase Gram-positive methylotrophs, in addition to the cyclic route (see below), employ mycothiol (1-O-(20 -[N-acetyl-L-cysteinyl]amido-dependent 20 -deoxy-a-D-glucopyranosyl)-D-myoinositol) FaDHs (Vorholt 2002). This trimeric enzyme consists of a single type of subunit containing Zn (Van Ophem et al. 1992). In addition, dye-linked enzymes have been described for some Gram-negative methylotrophs (Klein et al. 1994; Zahn et al. 2001), but their role in methylotrophy remains poorly understood.

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). 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). Typically, species employing this pathway do not contain genes or activities for the H4MPT-linked pathway. With this respect, it is interesting to point out that heterologously expressed genes for this pathway were able to complement mutants of M. extorquens AM1 deficient in genes of the H4MPT-linked pathway (Marx et al. 2003a).

NAD-Linked Formaldehyde Dehydrogenase A single enzyme system such as NAD-linked, glutathioneindependent FaDHs (exemplified by the well-studied enzyme from Pseudomonas putida; Tanaka et al. 2003) is not found in most methylotrophs. However, they appear to be the major formaldehyde detoxification systems in organisms such as Pseudomonas and non-methylotrophic Burkholderia species that, like methylotrophs, tend to possess multiple formaldehyde oxidation systems (Marx et al. 2004; Roca et al. 2009). Likely, this enzyme is involved in formaldehyde oxidation/detoxification in the methylotrophic Pseudomonas and Burkholderia species. However, no mutant tests have been performed yet with these organisms.

Aerobic Methylotrophic Prokaryotes

Cyclic Formaldehyde Oxidation Pathway Another formaldehyde oxidation 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. 7.3). The enzymes carrying out these reactions are those of the ribulose monophosphate (RuMP) cycle for formaldehyde assimilation, involving one additional enzyme, a 6-phosphogluconate dehydrogenase (Gnd). A glucose-6phosphate dehydrogenase (Zwf) is also required for this cycle, but this may or may not be a part of the assimilatory RuMP cycle depending upon the variant utilized (see below). While genes for Zwf are found in all methylotrophs and across life in general, and these show significant degree of conservation, two alternative Gnd genes have been recognized: GndA and GndB. The former is specific to NAD, and the latter is specific to NADP (Chistoserdova 2011). Methylotrophs that rely on the dissimilatory RuMP cycle for formaldehyde oxidation may employ either MtdA or MtdB (Lapidus et al. 2011), and some employ both. In the case of Methylobacillus flagellatus that encodes both enzymes, GndA was demonstrated to be of more importance than GndB in the organism’s fitness (Chistoserdova et al. 2000; Hendrickson et al. 2010). Glucose-6-phosphate dehydrogenase (Zwf) typically utilizes both NADP and NAD (Kiriuchin et al. 1988).

Distribution of Formaldehyde Oxidation Pathways in Methylotrophs Although some methylotrophs appear to have only one formaldehyde oxidation pathway (Giovannoni et al. 2008; Halsey et al. 2012), others have multiple routes, but their roles in methylotrophy are not necessarily redundant (Chistoserdova et al. 2009). One of the most persistent formaldehyde oxidation Hexulose 6-Phosphate CH2O

2

1

Fructose 6-Phosphate Ribulose 5-Phosphate

3

5 Glucose 6-Phosphate

2H 2H CO2

4

6-Phosphogluconate

. Fig. 7.3 Cyclic pathway of formaldehyde oxidation, involving enzymes of the RuMP pathway. 1 hexulosephosphate synthase, 2 hexulosephosphate isomerase, 3 fructosephosphate isomerase, 4 NAD(P)-dependent glucose 6-phosphate dehydrogenase, 5 NAD- or NADP-linked 6-phosphogluconate dehydrogenase (Adapted from Anthony (1982))

7

pathways is the pathway involving H4MPT-linked C1 derivatives. The only functional groups lacking this pathway are the minimalist marine methylotrophs possessing very small genomes, the autotrophic methylotrophs such as Paracoccus denitrificans, the methanotrophic Verrucomicrobia, and Gram-positive methylotrophs (Chistoserdova 2011). While in the serine cycle organisms, such as Methylobacterium species, the pathway has been found to be absolutely essential for methylotrophy capability (Chistoserdova et al. 2009), in Betaproteobacteria, it appears to play an auxiliary function (Chistoserdova 2011). While the pathway is present in all proteobacterial methanotrophs and the respective enzymes are present at high activities (Vorholt et al. 1999), its role in these organisms has not yet been investigated and other enzymes have been proposed to function in formaldehyde oxidation (Zahn et al. 2001). The analogous pathway linked to H4F and involving FolD bifunctional enzyme has been shown to operate as the main dissimilatory pathway during growth on C1 compounds whose degradation involves methyltransferase reactions resulting in formation of methyl-H4F (Studer et al. 2002). The role of this pathway in oxidizing formaldehyde has also been proposed for some organisms, based on the lack of any other recognizable formaldehyde oxidizing pathways in their genomes (Hou et al. 2008; Giovannoni et al. 2008). However, this would involve spontaneous condensation of formaldehyde with H4F (Chistoserdova 2011) physiological significance of which has been recently revised (Crowther et al. 2008). Thus, this proposal remains controversial (Chistoserdova 2011). The alternative variant of H4F-linked C1 transfer pathway, involving MtdA and Fch, is encoded in the genomes of methylotrophs possessing the complete or partial serine cycle (Chistoserdova 2011), and this pathway has been shown to operate in reductive rather than oxidative direction (Crowther et al. 2008; Chistoserdova et al. 2009). In Gram-positive methylotrophs, the mycothiol-linked FaDH has been proposed to constitute the main formaldehyde oxidation pathway (Duine 1999), and the GSH-linked formaldehyde oxidation system has been found to constitute the main formaldehyde oxidation pathway in autotrophic methylotrophs not possessing the H4MPT pathway (Harms et al. 1996; Barber and Donohue 1998). The cyclic formaldehyde oxidation pathway appears to be the main formaldehyde oxidative pathway in the methylotrophs utilizing the RuMP cycle (Anthony 1982; Hendrickson et al. 2010), while the H4MPT pathway plays an auxiliary role (Chistoserdova et al. 2000; Hendrickson et al. 2010). Additional formaldehyde detoxification systems may also be employed by these methylotrophs (Chistoserdova et al. 1991). The role of the dissimilatory RuMP cycle in gammaproteobacterial methanotrophs has been questioned based on the low activities of the glucose-6-phosphate and 6-phosphogluconate dehydrogenases (Zatman 1981).

Formate Oxidation Methylotrophs, like other organisms, possess at least one formate dehydrogenase (FDH), and most encode multiple FDH

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enzymes. For example, M. extorquens species possess four different FDH enzymes, which include a tungsten-containing NAD-linked FDH (FDH1; Laukel et al. 2003), a predicted molybdenum-containing NAD-linked FDH (FDH2; Chistoserdova et al. 2004b), a predicted cytochrome-linked FDH that is likely periplasmic (FDH3; Chistoserdova et al. 2004b), and a novel type of FDH whose electron acceptor remains unknown (FDH4; Chistoserdova et al. 2007a), and all of these appear to be functional. However, when each of the enzymes was disrupted in a wild-type background, mutants in only one of them, FDH4, revealed a phenotype: These mutants accumulated formate in the late stationary phase of growth on methanol (Chistoserdova et al. 2007a). If an intact FDH4 was present, the three other enzymes appeared redundant during growth on methanol. However, the situation was different when the organism grew on formate. FDH4 alone could not sustain the growth, and at least one of the other FDH enzymes was needed (Chistoserdova et al. 2004b). FDH4 in this organism also appeared to be somehow involved in acid stress response, but the mechanism of this involvement remains unknown (Chistoserdova et al. 2007a). In organisms oxidizing formaldehyde via the RuMP cycle, the formate oxidation step is expected to be less critical for methylotrophy, and, accordingly, activities of FDH enzymes are typically measured at very low levels (Anthony 1982). Relative contributions of alternative FDHs to methylotrophy were recently investigated in a RuMP cycle methylotroph, Methylobacillus flagellatus that possesses FDH enzymes homologous to FDH2 and FDH4 of M. extorquens (Hendrickson et al. 2010). It was concluded from this study that the formate oxidation step is essential even when the dissimilatory RuMP cycle is functionally present, as no mutants could be generated simultaneously lacking both FDHs, and FDH4 appeared to play a more important function in fitness of this organism compared to FDH2 (Hendrickson et al. 2010). Overall, based on the available genomic data, FDHs are not conserved among methylotroph groups, but rather these are randomly distributed. For example, close homologs of the FHD2 enzymes described above are widespread across methylotrophic and non-methylotrophic Proteobacteria, while non-proteobacterial methylotrophs encode FDH enzymes homologous to the ones of other representatives of respective (or related) phyla (Hou et al. 2008; Ettwig et al. 2010).

Assimilatory Metabolism Three main pathways of assimilatory metabolism are known in aerobic methylotrophs: the ribulose monophosphate cycle that assimilates carbon at the level of formaldehyde (Anthony 1982), the serine cycle that assimilates carbon at the levels of methylene-H4F and CO2 (1:1 ratio; Peyraud et al. 2009; Chistoserdova et al. 2009), and the Calvin–Benson–Bassham (CBB) cycle that assimilates carbon exclusively at the level of CO2

(Anthony 1982). In the two former pathways, 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. 7.4, 7.5).

Serine Cycle The serine cycle is one of the assimilatory pathways specific to methylotrophy. The pathway is initiated with the condensation of methylene-H4F and glycine to form serine. This 3-carbon compound then undergoes a series of transformations to phosphoenolpyruvate, which is carboxylated to form malate, followed by CoA derivatization by malate thiokinase. The resulting malyl-CoA is cleaved into two 2-carbon compounds, glyoxylate, and acetyl-CoA, the former being converted back into glycine via serine glyoxylate aminotransferase and the second undergoing series of transformations to regenerate the second molecule of glyoxylate, thus completing the cycle (> Fig. 7.4). In many methylotrophs with the serine cycle, acetyl-CoA is converted to glyoxylate via the recently described ethylmalonyl-CoA pathway, while in others, this conversion involves the more traditional isocitrate lyase shunt (see below). Serine cycle genes are typically found in clusters and are subjects of coordinated regulation (Kalyuzhnaya and Lidstrom 2003, 2005). Such clusters have been analyzed in a number of methylotrophs and potential methylotrophs (Vuilleumier et al. 2009), and their structure and content suggest a complex history. For example, the structure of the serine cycle gene cluster first characterized in M. extorquens AM1 (Chistoserdova et al. 2003) is highly conserved in some alphaproteobacterial methylotrophs, and these clusters are remarkably syntenic with the cluster in a betaproteobacterial methylotroph, Methylibium petroleiphilum (Vuilleumier et al. 2009). However, the genome of Ruegeria pomeroyi, also an Alphaproteobacterium, contains a cluster that shows no synteny to the M. extorquens cluster (Vuilleumier et al. 2009). In addition, some of the genes in the cluster reveal very low levels of homology to their counterparts in other Alphaproteobacteria, suggesting independent evolution of the serine cycle in different bacterial lineages. Gene clustering is also different in Hyphomicrobium denitrificans, and some of the genes (such as that encoding phosphoenolpyruvate carboxylase and hydroxypyruvate reductase) show little homology with other phosphoenolpyruvate carboxylase and hydroxypyruvate reductase genes (Chistoserdova 2011). Genomes of gammaproteobacterial methanotrophs also harbor most of the serine cycle genes, but few of them are clustered (Ward et al. 2004). However, the gene for one key enzyme of the serine cycle, the phosphoenolpyruvate carboxylase, is missing from the genomes. Thus, the cycle might be incomplete in these bacteria, or a modification of the cycle using an alternative enzyme is operational. The proteomic analysis of Methylococcus capsulatus revealed that most of the serine cycle genes are expressed during growth on methane (Kao et al. 2004).

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Aerobic Methylotrophic Prokaryotes 2 2-Phosphoglycerate to Biomass

S5

E10

CO2 3 2-Phosphoglycerate

1 Phosphoenolpyruvate

E11

Succinyl-CoA Methylmalonyl-CoA

CO2

S6 Succinate

S4

1 Oxaloacetate

E12

S8/E8

Fumarate

3 Glycerate

E9

Propionyl-CoA β-Methylmalyl-CoA

S7 E13

E7

S3 2 Malate

Serine cycle

3 Hydroxypyruvate

Mesaconyl-CoA

S8/E8

E6

2 Malyl-CoA

S2

S9/E14 3 Glyoxylate 3 Serine

3 CH2=H4F

S1

E5 EthylmalonylCoA

2 Acetyl-CoA

E4

E1 S2

Methylsuccinyl-CoA

Ethylmalonyl-CoA Pathway

Crotonyl-CoA Acetoacetyl-CoA

3 Glycine

E2

CO2

E3

β-Hydroxybutyryl-CoA

3 CH2=H4F + 3 CO2= 2 2-Phosphoglycerate

. Fig. 7.4 The serine cycle for formaldehyde assimilation and the ethylmalonyl-CoA (EMC) pathway. Reactions of the serine cycle are denoted with letter ‘‘S’’; enzymes of the EMC pathway are denoted with letter ‘‘E,’’ and enzymes shared between the pathways are denoted with both letters. S1 serine hydroxymethyltransferase, S2 serine glyoxylate aminotransferase, S3 hydroxypyruvate reductase, S4 glycerate kinase, S5 enolase, S6 phosphoenolpyruvate carboxylase, S7 malate dehydrogenase, S8/E8 malate thiokinase, S9/E14 malyl-CoA/ b-methylmalyl-CoA lyase, E1 b-ketothiolase, E2 acetoacetyl-CoA reductase, E3 crotonase, E4 crotonyl-CoA reductase/carboxylase, E5 ethylmalonyl-CoA mutase, E6 methylsuccinyl-CoA dehydrogenase, E7 mesaconyl-CoA hydratase, E9 propionyl-CoA carboxylase, E10 methylmalonyl-CoA mutase, E11 succinyl-CoA dehydratase, E12 succinate dehydrogenase, E13 fumarase (Adapted from Chistoserdova et al. (2009))

Glyoxylate Regeneration The assimilation of C1 units via the serine cycle requires regeneration of glyoxylate from acetyl-CoA. It has been recognized early on that, while this task can be carried out by the classic glyoxylate shunt, many serine cycle methylotrophs employ instead an alternative glyoxylate regeneration pathway (Anthony 1982) that remained a puzzle for over three decades. This pathway has been recently resolved by a combination of efforts from different laboratories, and it is now known as the ethylmalonylCoA pathway (EMCP; Erb et al. 2007, 2008; Peyraud et al. 2009; Chistoserdova et al. 2009; > Fig. 7.4). A historical account describing half a century that was required to solve this pathway has been recently published (Anthony 2011). The pathway shares reactions and enzymes with the serine cycle (malate thiokinase, malyl-CoA lyase), the tricarboxylic acid cycle (succinate dehydrogenase, fumarase), the polyhydroxybutyrate cycle (b-ketothiolase, acetoacetyl-CoA reductase), and with other metabolic pathways (methylmalonyl-CoA mutase, propionylCoA carboxylase), in addition to the specific reactions such as

ethylmalonyl-CoA mutase (Erb et al. 2008) and crotonyl-CoA reductase/carboxylase (Erb et al. 2007). Respectively, genes for the latter enzymes serve as markers for probing genomes for the presence of the EMCP. The EMCP is not specific to methylotrophy. It is also utilized for metabolism of C2 compounds by both methylotrophs, such as M. extorquens (Okubo et al. 2010; Schneider et al. 2011), and non-methylotrophic bacteria such as Rhodobacter and Streptomyces (Alber 2010). In the latter case, genes for the EMCP are present in the genomes but not the genes for the serine cycle. Some methylotrophs, however, opt to use the glyoxylate shunt instead of the EMCP. For example, the genomes of Methylibium petroleiphilum and Methylocella silvestris, while lacking genes for the key enzymes of the EMCP, contain the genes for the glyoxylate shunt (Kane et al. 2007; Chen et al. 2010a), and the genome of Hyphomicrobium denitrificans encodes both pathways (Chistoserdova 2011). In the genome of M. capsulatus, neither the EMCP nor the glyoxylate shunt appears to be encoded, consistent with the hypothesis of the serine pathway fulfilling an auxiliary metabolic task (Chistoserdova et al. 2005b).

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3 CH2O Fructose 6-Phosphate 3 Ribulose 5Phosphate

3 Hexulose 6Phosphate

Fructose 6Phosphate ATP

Ribose 5Phosphate

Glyceraldehyde 3Phosphate

Fructose 6Phosphate

or

6-Phosphogluconate Fructose 1,6Bisphosphate

Erythrose 4Phosphate Xylulose 5- Sedoheptulose 7Phosphate Phosphate Xylulose 5Phosphate

ADP

Fructose 6Phosphate

Glucose 6-Phosphate NAD(P) NAD(P)H+H+

Glyceraldehyde Dihydroxyacetone 3-Phosphate Phosphate

2-Keto 3-Deoxy 6Phosphogluconate

Glyceraldehyde Pyruvate 3-Phosphate

or Fructose 6-Phosphate ATP ADP Fructose 1,6-Bisphosphate

Glyceraldehyde 3-Phosphate

Sedoheptulose 7Phosphate

Dihydroxyacetone Phosphate Erythrose 4Phosphate Sedoheptulose 1,7Bisphosphate

Pi

. Fig. 7.5 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))

Ribulose Monophosphate Cycle The ribulose monophosphate cycle (RuMP cycle) is shown in > Fig. 7.5. Formaldehyde is condensed with the acceptor molecule (RuMP) by the enzyme hexulose phosphate synthase (Hps) to produce hexulose phosphate. The six-carbon molecule is then isomerized to fructose 6-phosphate by phosphohexose isomerase (Hpi), and a series of interconversions occur that regenerate the five-carbon acceptor molecule. The condensation of three formaldehyde molecules results in the net production of one C3 compound (Anthony 1982). Theoretically, four different variants of the ribulose monophosphate pathway are possible, representing different combinations of two alternative sugar cleavage scenarios (cleavage of fructose bisphosphate, via fructose bisphosphate aldolase versus cleavage of 2-keto 3-deoxy 6-phosphogluconate, via a specific aldolase) and two alternative formaldehyde acceptor (RuMP) regeneration scenarios (one involving transaldolase and one involving sedoheptulose 1,7-bisphosphatase; Anthony 1982).

It is worth noting that the presence of the hps/hpi gene pair is not always a sign of methylotrophy capability as the same cycle is utilized by some non-methylotrophs, serving as an additional means for formaldehyde detoxification (Yasueda et al. 1999). Indeed, homologs of hps/hpi are found in the genomes of a variety of non-methylotrophic species, including Bacillus, Escherichia, etc., and also in some Archaea (Kato et al. 2006).

The CBB Cycle A number of methylotrophs possess genes and enzymes of the CBB cycle, some as the only means for C1 assimilation and others in addition to other C1 assimilatory cycles. In the latter case, the contribution of the CBB cycle to methylotrophy remains poorly understood (Baxter et al. 2002; Chistoserdova et al. 2005b). The three major groups that rely on the CBB for methylotrophy are (1) the alphaproteobacterial autotrophs

Aerobic Methylotrophic Prokaryotes

(such as Paracoccus denitrificans and Xanthobacter autotrophicus; Baker et al. 1998), (2) methanotrophic Verrucomicrobia (Methylacidiphilum infernorum; Op den Camp et al. 2009; Khadem et al. 2011), and (3) methanotrophs belonging to the NC10 phylum (Ettwig et al. 2010). Based on sequence comparisons, the key CBB genes (cbbLS) may be subjects of lateral transfers. For example, two cbb gene clusters are found in the genome of Methylibium petroleiphilum, one that is more related to the clusters in other Betaproteobacteria, the other more related to the clusters in Alphaproteobacteria (Kane et al. 2007). While analysis of the genome of the NC10 representative Candidatus Methylomirabilis oxyfera revealed that most of the C1 metabolism genes significantly diverge from the genes found in other groups of methylotrophs, the cbb operon showed high homology with the operons from Proteobacteria (Ettwig et al. 2010). The cbb genes of M. acidiphilum are most closely related to the genes in Chloroflexi and Actinobacteria.

Methylotrophic Bacteria Methanotrophs Methanotrophs are the subgroup of the methylotrophic bacteria that 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; Pol et al. 2007; Dunfield et al. 2007; Islam et al. 2008; Antony et al. 2010). They contain characteristic intracytoplasmic membrane systems (Hanson and Hanson 1996; Trotsenko and Murrell 2008), either stacks of membrane disks in the type I strains (Gammaproteobacteria), rings of membranes at the periphery of the cell in the type II strains (Methylosinus and Methylocystis), vesicular membranes in Methylocella (Dedysh et al. 2000), a single-membrane stack parallel to the cell wall in Methylocapsa (Dedysh et al. 2002), or carboxysome-like structures or vesicular membranes in Verrucomicrobia (Op den Camp et al. 2009). Until recently, all well-studied methanotrophs have been assumed to be obligate methylotrophs, unable to grow on compounds with C–C bonds. However, it has now been reported that some methanotrophs, including representatives of the classic methanotroph guilds such as Methylocystis, Methylosinus, and Methylocella are capable of growth on a range of multicarbon compounds (Semrau et al. 2011), and this capability has been proposed to be significant for environmental fitness of these strains (Dunfield et al. 2010). 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; Trotsenko and Murrell 2008). Alphaproteobacterial methanotrophs utilize the serine cycle for C1 assimilation, and some of them, specifically representatives of the family Beijerinckiaceae, also contain the

7

genes of the CBB cycle. Gammaproteobacterial methanotrophs primarily utilize the RuMP cycle as their major assimilatory pathway. However, some of them also encode elements of the serine cycle and some encode reactions of the CBB cycle. Moreover, the latter two pathways were shown to be expressed, along with the RuMP cycle, suggesting that they may play a role in C1 assimilation (Baxter et al. 2002; Kao et al. 2004). The recently discovered methanotrophs belonging to Verrucomicrobia and methanotrophs of the NC10 phylum do not contain recognizable genes for either the serine or the RuMP cycles. Thus, they appear to be truly autotrophic methanotrophs (Op den Camp et al. 2009; Khadem et al. 2011; Ettwig et al. 2010) (> Table 7.1). Methanotrophs exist as symbionts in mussels, clams, and Pogonophora, and although the 16 S rRNA sequences class with type I methanotrophs, they have not yet been isolated in pure culture (Distel and Cavanaugh 1994). Symbiosis of methanotrophs was also demonstrated with Sphagnum plants (Raghoebarsing et al. 2005; Kip et al. 2010).

Non-methane-Utilizing Methylotrophs The bacteria capable of growth on methanol and other methylated compounds but not on methane are also diverse and are widely distributed in terrestrial, freshwater, and marine habitats (Anthony 1982). Bacteria capable of utilizing methylated amines are particularly prevalent in the marine environment (Neufeld et al. 2008a; 2008b) where it is postulated that they may play a role in carbon cycling in the photic zone (Strand and Lidstrom 1984). The pink-pigmented Methylobacterium strains are common epiphytes on plant leaves, and some evidence exists to suggest mutualistic relationships with plants (Holland and Polacco 1994; Sy et al. 2005; Knief et al. 2010). So far, all of the Gram-positive and alphaproteobacterial strains are facultative methylotrophs, whereas the betaproteobacterial and gammaproteobacterial strains are either obligate methylotrophs or restricted facultative methylotrophs (capable of growth on a restricted range of multicarbon compounds; Anthony 1982; Chistoserdova et al. 2009; Kalyuzhnaya et al. 2011). Many but not all of these bacteria can grow on methanol, and many will grow on other methylated compounds (Anthony 1982). The non-methane-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).

Genetics in Aerobic Methylotrophs Genetic Capabilities Some methylotrophs, most notably Methylobacterium extorquens AM1, have been established as model systems for genetic manipulations, with a tool box including controlled

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expression vectors (Marx and Lidstrom 2001), promoter probe vectors with a choice of reporters (Xu et al. 1993; Marx and Lidstrom 2001, 2004), suicide vectors for mutant generation (Chistoserdov et al. 1994; Marx and Lidstrom 2002), including systems for generating unmarked mutations (Marx and Lidstrom 2002; Marx 2008), and chromosomal insertion vectors (Marx and Lidstrom 2004). Using these tools, over a hundred mutations were generated in M. extorquens AM1 and multiple chromosomal regions were expressed, in order to address questions of specific roles for specific genes and pathways (Chistoserdova et al. 2003; Marx and Lidstrom 2004; Chistoserdova et al. 2007a). While so far at a lesser scale, similar tools have been employed in other Gram-negative methylotrophs (Ali et al. 2006; Ali and Murrell 2009; Crombie and Murrell 2011; Ojala et al. 2011). Tools have also been reported for genetic manipulations in Gram-positive methylotrophs (Nesvera et al. 1994; Vrijbloed et al. 1995; Cue et al. 1997).

Methylotroph Genomics The first genomic sequence for a methylotroph (a draft) that became publically available was for the model methylotroph, M. extorquens AM1 (Chistoserdova et al. 2003). This was quickly followed by the complete genome sequence of another model methylotroph, Methylococcus capsulatus Bath (Ward et al. 2004). In the following years, complete or high-quality draft genomic sequences of other major model methylotrophs became available (Chistoserdova et al. 2007a; Vuilleumier et al. 2009; Chen et al. 2010a; Stein et al. 2010, 2011; Boden et al. 2011a, b; Lapidus et al. 2011; Heggeset et al. 2012; Marx et al., 2012), as well as the sequences of the newly-described organisms, including novel types of methanotrophs (Hou et al. 2008). Most of the phyla known for the methylotrophic lifestyle are now represented by genomic sequences, including gammaproteobacterial methanotrophs of the family Methylococcaceae (Ward et al. 2004; Boden et al. 2011; Svenning et al. 2011; Vuilleumier et al. 2012), alphaproteobacterial methanotrophs of the families Methylocystaceae and Beijerinckiaceae (Chen et al. 2010a; Stein et al. 2010, 2011), methanotroph species classified as Verrucomicrobia (Methylacidiphilaceae; Hou et al. 2008), and non-methanotrophic organisms belonging to Alpha-, Beta-, and Gammaproteobacteria as well as Gram-positive bacteria (> Table 7.1). In addition, genomes of some methylotrophs not available in pure cultures have been reconstructed from metagenomic data (Kalyuzhnaya et al. 2008b; Ettwig et al. 2010). The availability of a whole genomic sequence (metabolic blueprint) is becoming a prerequisite for informed analysis of metabolism of a given organism, at the same time providing the necessary basis enabling the downstream analyses such as transcriptomics, proteomics, and metabolomics that are parts of the systems biology approach to characterizing biological systems. M. extorquens AM1 became the first methylotroph model for system-approach-type investigations

(Skovran et al. 2010; Peyraud et al. 2011). Systems approaches for defining metabolic networks in other model methylotrophs are surely to follow in the near future.

Summary This chapter describes biochemical pathways operating in aerobic methylotrophic bacteria. We first define aerobic methylotrophy as a specific metabolic capability and describe the phylogenetic diversity of these bacteria. We then describe enzymes involved in primary oxidation of different singlecarbon substrates, resulting in formaldehyde, methyl-, or methylene radical, and describe the variety of pathways used for their assimilation and dissimilation. We also give a brief account of genetic manipulation tools in methylotrophic bacteria and examples of system approaches for studying their metabolism, including availability of whole-genome sequence information.

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8 Dissimilatory Fe(III)- and Mn(IV)-Reducing Prokaryotes Derek Lovley Department of Microbiology, University of Massachusetts, Amherst, MA, USA

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 Significance of Fe(III)- and Mn(IV)-Reducing Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 288 Oxidation of Organic Matter in Anaerobic Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 288 Influence on Metal and Nutrient Geochemistry and Water Quality . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289 Bioremediation of Organic and Metal Contaminants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289 A Possible Early Form of Microbial Respiration . . . . 290 Fe(III)- and Mn(IV)-Reducing Microorganisms Available in Pure Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290 Fermentative Fe(III)- and Mn(IV)-Reducing Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290 Sulfate-Reducing Microorganisms . . . . . . . . . . . . . . . . . . 290 Microorganisms That Conserve Energy to Support Growth from Fe(III) and Mn(IV) Reduction . . . . . . . . . . . 292 Geobacteraceae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 292 Geothrix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 292 Geovibrio ferrireducens and Deferribacter thermophilus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 296 Ferribacter limneticum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 297 Shewanella–Ferrimonas–Aeromonas . . . . . . . . . . . . . . . . . 298 Sulfurospirillum barnesii . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298 Acidophilic Fe(III)-Reducing Microorganisms . . . . . 298 Hyperthermophilic and Thermophilic Archaea and Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298 Forms of Fe(III) and Mn(IV) That Can Serve as Electron Acceptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 299 Products of Fe(III) and Mn(IV) Reduction . . . . . . . . . 299 Mechanisms for Electron Transfer to Fe(III) and Mn(IV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 Other Respiratory Capabilities of FMR . . . . . . . . . . . . . . . . . 301 Electron Transfer to Other Metals and Metalloids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 301 Electron Transfer to and from Humic Substances and Other Extracellular Quinones . . . . . . . . . . . . . . . . . . 301 Proton Reduction in Syntrophic Association with Hydrogen-Consuming Microorganisms . . . . . . . . . . . . 301 Reductive Dechlorination . . . . . . . . . . . . . . . . . . . . . . . . . . . 302 Recovery of Fe(III)- and Mn(IV)-Reducing Microorganisms in Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 302 Localizing Zones of Fe(III) and Mn(IV) Reduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 302

Isolation Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 303 Suggested Media for Enrichment and Culturing of FMR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 304 Preparation of Fe(III) and Mn(IV) Forms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 305

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.

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_69, # Springer-Verlag Berlin Heidelberg 2013

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

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, 1991a, 1995a; Lovley et al. 1993a, 1997c; Nealson and Saffarini 1994). Therefore only highlights of the significance of Fe(III)reducing microorganisms, abstracted from these reviews, will be briefly summarized here.

Fe(III) reduction is the predominant mechanism for organic matter oxidation (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, 1995b). A model for the oxidation of organic matter in sedimentary environments in which Fe(III) reduction is the predominant terminal electron-accepting 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. 8.1), 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 longchain 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

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, 1995b; Canfield et al. 1993; 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

Aromatic Compounds

Ge

ob

ac

ter

ma

tal

ia phil

ac

lla s

ane hew

ro cha

S

Hydrolysis of Complex Organic Matter

Fermentable Substrates

Fermentative Microorganisms

Long Chain Fatty Acids

ler

ed uc en Acetate Ge oba s c o vib ter Minor ria , G fermentation Fe eot rri hri acids ac x ier Ge

rix Geoth acter, b o e G anella H2 Shew ns s nta tati rmo patmi e f rix cnas oth Ge furom ul Dex

Fe(II) CO2

. Fig. 8.1 Proposed pathways for organic matter degradation in mesophilic environments in which Fe(III) reduction is the predominant terminal electron-accepting process

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

TEAP. This 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 red-yellow 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 results 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 groundwater sources of drinking water, being one of the most prevalent groundwater quality problems (Anderson and Lovley 1997). 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, 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

8

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 Phillips 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 et al. 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 petroleumcontaminated aquifers (Lovley et al. 1989b; Lovley 1995c, 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 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, 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.

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

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 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 suggest that the last common ancestor was a hydrogenoxidizing, 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 quantities 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 8.1). 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 without 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 8.1]. 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) (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 cannot 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

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

8

. Table 8.1 Organisms known to reduce Fe(III) but not known to conserve energy from Fe(III) reduction Electron donor

Form of Fe(III) reduceda

Reference

Actinomucor repens

Glucose

Hematite

Ottow and von Klopotek (1969)

Aerobacter aerogenes

Glucose–asparagine

Hematite

Ottow (1970)

Aerobacter sp.

Glucose

PCIO

Bromfield (1954)

Alternaria tenuis

Glucose

Hematite

Ottow and von Klopotek (1969)

Bacillus cereus

Glucose–asparagine

Hematite

Ottow (1970)

Bacillus circulans

Sucrose

PCIO

Bromfield (1954)

Glucose–asparagine

Hematite

Ottow (1970)

Organism Fermentative bacteria

Sucrose

Ferromanganese ore

Troshanov (1968)

Bacillus mesentericus

Sucrose

Ferromanganese ore

Troshanov (1968)

Bacillus polymyxa

Glucose

PCIO

Roberts (1947)

Glucose

Hematite

Hammann and Ottow (1974)

Sucrose

PCIO

Bromfield (1954)

Glucose–asparagine

Hematite

Ottow (1970)

Bacillus pumilus Bacillus sp.

Glucose

Limonite, goethite, hematite

De Castro and Ehrlich (1970)

Bacillus subtilis

Glucose

Hematite

Ottow and Glathe (1971)

Bacteroides hypermegas

Glucose–tryptone

Fe(III)-Cl3

Jones et al. (1984a)

Clostridium butyricum

Glucose

Hematite

Hammann and Ottow (1974)

Clostridium polymyxa

Sucrose

Ferromanganese ore

Troshanov (1968)

Clostridium saccarobutyricum

Glucose

Hematite

Hammann and Ottow (1974)

Clostridium sporogenes

Glucose, peptone

PCIO

Starkey and Halvorson (1927)

Escherichia coli

Glucose, peptone

PCIO

Starkey and Halvorson (1927)

Escherichia coli

Glucose–asparagine

Hematite

Ottow (1970)

Fusarium oxysporum

Glucose

Ferric ammonium citrate

Gunner and Alexander (1964)

Fusarium oxysporum

Glucose

Hematite

Ottow and von Klopotek (1969)

Fusarium solani

Glucose

Hematite

Ottow and von Klopotek (1969)

Paracolobactrum sp.

Glucose

PCIO

Bromfield (1954)

Pseudomonas aeruginosa

Glucose–asparagine

Hematite

Ottow (1970)

Pseudomonas denitrificans

Glucose

Fe(III)-Cl3

Jones et al. (1984a)

Pseudomonas liquefaciens

Sucrose

Ferromanganese ore

Troshanov (1968)

Pseudomonas (several species)

Glucose–asparagine

Hematite

Ottow and Glathe (1971)

Rhodobacter capsulatus

Malate

Fe(III)-NTA

Dobbin et al. (1996)

Serratia marcescans

Glucose–asparagine

Hematite

Ottow (1970)

Sulfolobus acidocaldarius

Elemental sulfur

Fe(III)-Cl3

Brock and Gustafson (1976)

Thiobacillus thiooxidans

Elemental sulfur

Fe(III)-Cl3

Brock and Gustafson (1976) Kino and Usami (1982)

Vibrio sp.

Glucose

Fe(III)-Cl3

Jones et al. (1983)

Vibrio sp.

Malate, pyruvate

Fe(III)-Cl3

Jones et al. (1984b)

Wolinella succinogenes

Formate

Fe(III)-Cit

Lovley et al. (1998)

Acetate

Fe(III)-NTA

Lovley et al. (1993)

Sulfate-reducing bacteria Desulfobacter postgatei Desulfobacterium autotrophicum

H2

Fe(III)-NTA

Lovley et al. (1993)

Desulfobulbus propionicus

Propionate

Fe(III)-NTA

Lovley et al. (1993)

Desulfovibrio baarsii

Butyrate, caproate, octanoate

Fe(III)-NTA

Lovley et al. (1993)

Desulfovibrio desulfuricans

Lactate

Fe(III)-Cl3

Jones et al. (1984a) Coleman et al. (1993)

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292

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

. Table 8.1 (continued) Organism

Electron donor

Form of Fe(III) reduceda

Reference

Desulfovibrio baculatus

Lactate

Fe(III)-NTA

Lovley et al. (1993)

Desulfovibrio sulfodismutans

Lactate

Fe(III)-NTA

Lovley et al. (1993)

Desulfovibrio vulgaris

Lactate

Fe(III)-NTA

Lovley et al. (1993)

Desulfotomaculum nigrificans

Lactate

Fe(III)-Cl3

Jones et al. (1984a)

Archaeoglobus fulgidus

H2

Fe(III)-Cit

Vargas et al. (1998)

Methanococcus thermolithotrophicus

H2

Fe(III)-NTA

Vargas et al. (1998)

Methanopyrus kandleri

H2

Fe(III)-Cit

Vargas et al. (1998)

Pyrococcus furiosus

H2

Fe(III)-Cit

Vargas et al. (1998)

Pyrodictium abyssi

H2

Fe(III)-Cit

Vargas et al. (1998)

Archaea

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]

‘‘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) was tenfold 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 2,500-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 8.2) are phylogenetically (> Fig. 8.2) and morphologically (> Fig. 8.3) 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. 8.2; > Table 8.2). 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 8.2). Several species of Geobacter have the ability to anaerobically oxidize aromatic compounds, including the hydrocarbon toluene. Geobacteraceae are the Fe(III) reducers most commonly recovered from a variety of sedimentary environments when the culture media contain acetate as the electron donor and Fe(III) oxide or the humic acid analog anthraquinone-2,6disulfonate (AQDS) as the electron acceptor (Coates et al. 1996, 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 (Rooney-Varga et al. 1999; Synoeyenbos-West et al. 1999).

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

Complete ND

Ac, BtOH, EtOH Prop, Pyr Ac, Pyr

North Sea oil field

Anoxic muds

Marine sediments

Freshwater sediments

Marine sediments

Marine and freshwater muds

Desulfuromonas acetexigens

Desulfuromonas acetoxidans

Desulfuromonas chloroethenica

Desulfuromonas palmitatis

Desulfuromusa bakii

Ac

Marine sediments

Mine-impacted lake sediments

Marine sediments

Freshwater ditch

Freshwater ditch

Deep subsurface

Aquatic sediments

Desulfuromusa succinoxidans

Ferribacterium limneticum

Ferrimonas balearica

‘‘Geobacter akaganeitreducens’’

‘‘Geobacter arculus’’

‘‘Geobacter chapellei’’ (strain 172)

‘‘Geobacter grbicium’’ (strain TACP-2)

ND

Complete

Complete

Complete

Complete

Complete

Complete

Ac, Buty, EtOH, For, Prop, Pyr

Ac, EtOH For, Lac

Ac, BtOH, Buty, Bzo, EtOH, For, Fum, H2, Lac, Mal, Prop, PrOH, Pyr, Succ

Complete

Complete

ND

Ac, EtOH, For, Fum, H2, Mal, ND Prop, PrOH, Pyr, Succ

Lac

Ac

Ac

Desulfuromusa kysingii Freshwater anoxic muds

Ac

Ac, Fum, Lac, Lau, Pal, Ste, Succ

Ac

Ac, CAA, H2, Lac, Mal, Pept, NDf Pyr, Succ, Try, Valr, YE

Incomplete

For, Lac

Deep subsurface

Deferribacter thermophilus

Complete

21–31

PCE, TCE, Fum, S2

PCIO, Fe(III)-Cit

PCIO, Fe-NTA,

PCIO

PCIO, akaganeite

PCIO, Fe(III)-Cit

PCIO, Fe(III)-P

Fe(III)-NTA

Fe(III)-Cit, Fe(III)NTA

Fe(III)-NTA

30

S , Mal, Fum

30

Mn(IV), S0, Fum, Mal

AQDS

30

25

30

Mn(IV), So, Fum, Mal

Mn(IV), AQDS, Fum

37

Mn(IV), nitrate

25

30

S , Mal, Fum, DMSO, nitrate

Nitrate, Fum

25

S , Mal, Fum

40

30

30

Mn(VI), S , polysulfides, Fum, Mal Mn(IV), Glut, Mal, Fum

60

60

37

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

Rod

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)

Lonergan et al. (1996)

Liesack and Finster (1994), Lonergan et al. (1996)

Lonergan et al. (1996)

Coates et al. (1995)

Krumholz (1997)

Roden and Lovley 1993

Coates et al. (1995)

Greene et al. (1997)

Boone et al. (1995)

Knight and Blakemore (1998)

Growth temp ( C) Morphology Referencee

Mn(VI), nitrate

Mn(IV), nitrate, TMAO

U(VI), Co(III), selenate, nitrate, Fum, O2

Other electron acceptorsd

Mn(IV), AQDS, S0, Fum PCIO, Fe(III)-Cit, Fe(III)-NTA, Fe(III)P

Fe(III)-NTA

Fe(III)-Cit, Fe(III)NTA

PCIO

PCIO, Fe(III)-Cit

Fe(III)-Cl3

PCIO, Fe(III)-Cit

Oxidation Fe forms with Fe(III)b reducedc

Bacillus infernus

Electron donors oxidized with Fe(III)a Glyc, Lac, Succ

Source

Aeromonas hydrophila Freshwater and sewage

Organism

. Table 8.2 Organisms known to conserve energy to support growth from Fe(III) reduction

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

8 293

Contaminated ditch Ac, For, H2

Contaminated aquifer

Geobacter sulfurreducens

Geothrix fermentans

Estuarine sediment

Aquatic sediments and other diverse environments

Shewanella alga

Shewanella putrefaciens

Incomplete Incomplete

H2, Lac For, H2, Lac Pyr

PCIO, Fe(III)-cit

PCIO, Fe(III)-cit

PCIO, Fe(III)-cit

ND

H2, Pept, YE

Marine hydrothermal waters

Pyrobaculum aerophilum

Rod Rod

100

30

Mn(VI), U(VI)*, S2O32 , AQDS, TMAO, Fum, O2

Mn(VI), U(VI), S , S2O32 , AQDS, 30 nitrate, Fum, O2

Rod

Rod

Rod

Rod

Rod

Rod

Nitrate, nitrite, O2

Mn(IV)*, U(VI)*, Co(III)*, Tc(VII)*, 100 Cr(VI)*, Au(III)*, Cyst, Glut, S0, SO32 , S2O32

PCIO, Fe(III)-cit

ND

H2, Pept, YE

Geothermal water

Pyrobaculum islandicum

Nitrate, O2

PCIO

H2

Swampy soil

‘‘Pseudomonas sp.’’

30

S

30

S

Fe(III)-NTA

30

S

Incomplete

EtOH, For, H2

Freshwater sediments

Pelobacter venetianus

Fe(III)-NTA

35

Co(III), S

Fe(III)-NTA

Lac

Pelobacter propionicus

Incomplete

Rod

30

Mn(IV), AQDS, S

Vibrio

Rod

Tc(VII)*, Co(III), AQDS, S , Fum, 35 Mal

Rod

Rod

30

30

Mn(IV), Tc(VII)*, U(VI), AQDS, humics, nitrate

AQDS, Fum

Rod

Myers and Nealson (1988), Lovley et al. (1989)

Caccavo et al. (1992)

Kashefi et al. 1999

Kashefi et al. 1999

Balashova and Zavarzin (1980)

Lonergan et al. (1996)

Lonergan et al. (1996)

Lovley et al. (1995)

Caccavo et al. (1996)

Coates et al., 1999

Caccavo et al. (1994)

Lovley and Philips 1988; Lovley et al. 1993

Coates et al. (1996)

Coates et al. (1998)

Growth temp ( C) Morphology Referencee

Mn(IV), AQDS S , nitrate, Fum, 30

Other electron acceptorsd

Incomplete

EtOH, H2

PCIO, Fe(III)citrate, Fe(III)-P

Geovibrio ferrireducens Contaminated ditch Ac, CAA, Fum, H2, Lac, Pro, Complete Prop, Pyr, Succ, YE

Pelobacter carbinolicus Marine sediments

PCIO, Fe(III)-Cit, Fe-NTA

PCIO, Fe(III)-Cit, Fe(III)-P

Complete

Ac, Lac

Complete

PCIO, Fe(III)-Cit

Ac, Bz, BzOH, BtOH, Buty, Bzo, BzOH, p-CR, EtOH, pHBz, p-HBzo, p-HBzOH, IsoB, IsoV, Ph, Prop, PrOH, Pyr, Tol, Valr

Complete

Aquatic sediments

Geobacter metallireducens

PCIO, Fe(III)-Cit

Ac, Buty, Bzo, EtOH, For, H2, Complete Prop, Pyr, Suc

Contaminated aquifer

‘‘Geobacter hydrogenophilus’’ (strain H2)

PCIO, Fe(III)-Cit

Oxidation Fe forms with Fe(III)b reducedc Complete

Ac, EtOH, For, H2, Lac

Contaminated wetland

‘‘Geobacter humireducens’’ (strain JW3)

Electron donors oxidized with Fe(III)a

Source

Organism

8

. Table 8.2 (continued)

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

Aquatic sediments

Freshwater marsh

Geothermally heated sea floor

Hot springs, Yellowstone

Deep gold-mine groundwater

Mine drainage

Shewanella saccharophilia

Sulfurospirillum barnesii

Thermotoga maritima

Thermoterrabacterium ferrireducens

Thermus strain SA-01

Thiobacillus ferrooxidans

Incomplete

For, Glyc, H2, Lac, Pyr, Suc, YE

ND ND

S0

Incomplete

H2, Glyc Lac

Fe(III)-Cit

ND

H2

Fe2(SO4)3

Fe(III)-Cit, Fe(III)NTA

PCIO, Fe(III)-Cit

PCIO, Fe(III)-cit

PCIO, Fe(III)-cit, Fe-NTA, Fe(III)-P, Fe(III)-EDTA

PCIO Fe(III)-cit

Incomplete

For, H2, Lac

Incomplete

For, EtOH, H2, Lac, Mal

S , O2,

Mn(VI), Co(III)*, Cr(VI)*, U(VI)*, S0, AQDS, nitrate, O2 30

65

65

AQDS, S2O32 , Fum

Rod

Rod

Rod

Rod

80

S0

Rod

Vibrio

30

Mn(IV), U(VI) *, S , AQDS, S2O32 , nitrate, Mal, Fum, O2

Rod

Mn(IV), selenate, arsenate, 30 S2O32 , S , nitrite, nitrate, Fum. TMAO, O2

30

Mn(IV), SO32 , nitrate, Fum, TmaO

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)

Boone personal communication

Abbreviations for electron donors and acceptors: acetate Ac, anthraquinone-2,6-disulfonic acid AQDS, alanine Ala, aspartate Asp, benzaldehyde Bz, benzoate Bzo, benzyl alcohol BzOH, 1,2 butanediol 1,2 Bu, butanol BtOH, butyrate Buty, casamino acids CAA, casein Cas, cystine Cyst, dimethyl sulfoxide 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, p-hydroxybenzyl alcohol 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) ethylenediaminetetraacetic 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 et al. 1999c Coates et al. Coates et al. 1999a Patrick et al. 1999

a

Subsurface

‘‘Shewanella putrefaciens CN32’’

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

8 295

296

8

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

0.1 substitutions/site

100

Desulfuromonas chloroethenica Desulfuromonas aetexigens

62 30

Pelobacter renetianus Pelobacter carbinolicus Desulfuromonas polmitatis 100

Desulfuromonas succinoxidans

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 94

Ferribacterium limneticum 97

87

Ferrimanas balearica Aeromonas hydrophila

100 100

Shewanella saccharophilia Shewanella putrefaciens Shewanella alga Geobacter fementans

100 97 100 100

Deferribacter themophilus Geovibria femreucens Bacillus infenus Themnoterrahacterium ferrireducens Themus sp.SA-01

Thenmotoga marthma 100

Pyrobaculum aerophilum Pyrobaculum islandicum

. Fig. 8.2 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 100 replicates

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 8.2), 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 petroleumcontaminated aquifer (Anderson et al. 1998). However, analyses of 16S rDNA sequences have indicated that Geothrix sp. is probably several orders of magnitude less numerous than

Geobacter species in such environments (Rooney-Varga et al. 1999; Synoeyenbos-West et al. 1999).

Geovibrio ferrireducens and Deferribacter thermophilus Culturing from hydrocarbon-impacted soils and a petroleum reservoir has 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

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

8

. Fig. 8.3 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

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,

297

298

8

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

it does not utilize Mn(IV) as an electron acceptor. To date, this organism has only been recovered from mining-impacted lake sediments.

Shewanella–Ferrimonas–Aeromonas In contrast to the organisms discussed above, which only grow anaerobically, several genera within the g-Proteobacteria can grow aerobically and, under anaerobic conditions, can use Fe(III), Mn(IV), or other electron acceptors (> Table 8.2). 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.

Sulfurospirillum barnesii Sulfurospirillum barnesii which was initially isolated based on its ability to use selenate as an electron acceptor (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 ferrooxidans 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 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).

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

As summarized in > Tables 8.1 and > 8.2, 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, 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 8.1). 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, 1986b; Phillips et al. 1993). The use of poorly crystalline Fe(III)-oxide as the Fe(III) form permitted the first recovery of 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 8.2; 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

8

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, 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)-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, 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 birnessite, a readily synthesized Mn(IV) oxide (see method for synthesis below). However, there is a wide diversity of Mn(IV) oxides 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 remain in solid forms (Lovley 1991a, 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 (Fe3PO4·8H2O),

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

and rhodochrosite (MnCO3; Lovley 1991a, 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 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. 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, 1998, 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) suggest 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, 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 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.

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

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 8.2). Fumarate is reduced to succinate, and S is 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 8.2]. 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 sulfate-reducing 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 8.2). 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 8.2). 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 Se and As(V) to As(III) (Laverman et al. 1995).

8

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 humic analog anthraquinone-2,6-disulfonate (AQDS) Lovley et al. 1996, 1998, 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 (Lovley et al. 1996, 1998; Coates 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 electronaccepting 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 make it preferable to carry out some studies on microbial reduction of extracellular quinones with humic analogs, such as AQDS (Lovley et al. 1996, 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 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

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

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 tetrachloroethylenerespiring 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 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 end products 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 also may 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 pore water 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 Phillips 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 [2-14 C]-acetate (Lovley 1997a). The reduction of Fe(III) can be considered to be the TEAP if (1) a tracer quantity of [2-14 C]-acetate added to the sediments is converted to 14CO2 with no production of 14 CH4, (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 14CH4 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 cannot be an important TEAP in the absence of nitrate, and (4) Mn(IV) reduction cannot 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 radiolabeled Fe(III) and other iron pools, including Fe(II). 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) over time. It is important that the solid-phase Fe(II) and Mn(II) pools be measured after acidic extractions or some other technique

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

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 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 8.2. An example of a freshwater and a marine medium is 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

8

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 these electron donors potentially could 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, nonmagnetic 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 humic 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) cannot reduce AQDS (Lovley et al. 1996, 1998, 2000). Recovery of AQDS-reducing microorganisms either through enrichment and isolation procedures or dilution-to-extinction approaches yields organisms that also can

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reduce iron [Fe(III); (Coates et al. 1998)]. The reduction of AQDS to AHQDS is 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-to-extinction 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.

RST Minerals Stock 50X Stock contains, per 100 ml NH4Cl

5.0 g

KCl

0.5 g

KH2PO4

0.5 g

MgSO4 · 7H2O

1.0 g

CaCl2 · 2H2O

0.1 g

Salt Stock Stock contains, per 100 ml MgCl2·6H2O

21.2 g

CaCl2·2H2O

3.04 g

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

Vitamin Solution Solution contains, per liter Biotin

2.0 mg

Folic acid

2.0 mg

Pyridoxine HCl

10.0 mg

Riboflavin

5.0 mg

Thiamine

5.0 mg

Nicotinic acid

5.0 mg

Pantothenic acid

5.0 mg

B-12

0.1 mg

NaHCO3

2.50 g

p-Aminobenzoic acid

5.0 mg

NH4Cl

0.25 g

Thioctic acid

5.0 mg

NaH2PO4·H2O

0.60 g

KCl

0.10 g

Vitamin solution

10.0 ml

Mineral solution

10.0 ml

Mineral Solution Solution contains, per liter

Bring solution to a final volume of 1 l. Media is dispensed, sparged with an 80:20 mixture of N2:CO2 gas, and then autoclaved.

Trisodium nitrilotriacetic acid

1.5 g

MgSO4

3.0 g

MnSO4·H2O

0.5 g

Marine Medium

NaCl

1.0 g

FeSO4·7H2O

0.1 g

Medium contains, per liter

CaCl2·2H2O

0.1 g

NaCl

20.0 g

CoCl2·6H2O

0.1 g

KCl

0.67 g

ZnCl2

0.13 g

NaHCO3

2.5.0 g

CuSO4 · 5H2O

0.01 g

Vitamin solution

10.0 ml

AlK(SO4)2·12H2O

0.01 g

Mineral solution

10.0 ml

H3BO3

0.01 g

RST minerals stock

20.0 ml

Na2MoO4

0.025 g

Salt stock∗

50.0 ml

NiCl2·6H2O

0.024 g

Na2WO4·2H2O

0.025 g

∗

Add salt solution aseptically and anaerobically after autoclaving.

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

Preparation of Fe(III) and Mn(IV) Forms Poorly Crystalline Fe(III) Oxide

Dissolve FeCl3·6H2O in water to provide final concentration of 0.4 M. 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 min 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 min. 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)-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 10 N NaOH. When the pH approaches 5.0, add the NaOH dropwise. Add medium constituents as outlined above. Bring to a final volume of 1 l. Do not expose this media to direct sunlight to prevent photoreduction of the 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 FeCl3·6H2O. 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).

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Manganese Oxide

To 1 l of a solution containing 80 mM NaOH and 20 mM KMnO4 slowly add 1 l 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-probable-number (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 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/l) 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 is the preferred method 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).

Goethite

Prepare a 0.4 M FeCl3·6H2O 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 h 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.’’

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 is also in

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its 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.

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9 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes Ralf Rabus1 . Theo A. Hansen2 . Friedrich Widdel3 1 Institute for Chemistry and Biology of the Marine Environment (ICBM), University of Oldenburg, Oldenburg, Germany 2 Microbial Physiology (MICFYS), University of Groningen, Groningen, The Netherlands 3 Max-Planck-Institut fu¨r Marine Mikrobiologie, Bremen, Germany

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310 Historical Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310 Sulfate-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310 Sulfate-Reducing Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 313 Sulfur-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 315 Sulfur-Reducing Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 316 Overview of Principal Properties . . . . . . . . . . . . . . . . . . . . . . . . . . 316 Sulfate-Reducing Bacteria and Archaea . . . . . . . . . . . . . . . . . 316 Sulfur-Reducing Bacteria and Archaea . . . . . . . . . . . . . . . . . . 321 Physiology, Biochemistry, and Molecular Biology . . . . . . . . . 322 Sulfate-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 322 Reduction of Sulfate to Sulfide . . . . . . . . . . . . . . . . . . . . . . 322 Dismutation of Sulfur Species . . . . . . . . . . . . . . . . . . . . . . . 333 Electron Acceptors Other than Sulfate . . . . . . . . . . . . . . 334 Electron Carriers and Possible Functions . . . . . . . . . . . 337 Metabolism of Electron Donors and Energy Conservation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339 Fermentative and Syntrophic Growth in the Absence of Sulfate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 361 Carbon Assimilation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 362 Assimilation of Nitrogen Compounds . . . . . . . . . . . . . . 363 Sulfate-Reducing Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Reduction of Sulfate to Sulfide . . . . . . . . . . . . . . . . . . . . . . 363 Electron Carriers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 Ferredoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 365 Sulfur-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . 367 Polysulfide Reductase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 368 Electron Transport from Formate or H2 to Polysulfide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 369 Regulation of Sulfur Respiration . . . . . . . . . . . . . . . . . . . . 370 Electron Acceptors Other than Sulfur . . . . . . . . . . . . . . . 370 Research on Desulfuromonas and Desulfurella . . . . . . 370 Sulfur-Reducing Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371 Reduction of Sulfur and Polysulfide . . . . . . . . . . . . . . . . . 372 Metabolism of Organic Electron Donors . . . . . . . . . . . 372 Autotrophic Carbon Assimilation . . . . . . . . . . . . . . . . . . . 373 Detoxification of Superoxide . . . . . . . . . . . . . . . . . . . . . . . . 373 Microorganisms Reducing Sulfur Compounds Other than Sulfate or Sulfur . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373

Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373 Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 Genetic Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 Sulfate-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 Genome Sizes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 374 Cloning, Sequencing, and Expression of Genes . . . . . 374 Physiological and Practical Prerequisites for Genetic Studies in Sulfate-Reducing Bacteria . . . . . . . 375 Delivery Systems for DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Transduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Conjugation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 Endogenous Plasmids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Transformation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Creation of Mutants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Chemical Mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Transposon Mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Gene Deletion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Site-Directed Mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Sulfur-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Genome Sizes, Genomic Libraries, and Cloning of Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 378 Physiological and Practical Prerequisites for Genetic Studies in Sulfur-Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . 379 Gene Deletion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379 Sulfur-Reducing Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 379 Abstract This chapter provides an overview of prokaryotes that reduce oxygenated sulfate or elemental sulfur in their energy metabolism. 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 ). Sulfur-reducing strains reduce elemental sulfur (or other lower oxidation states of this element, S0, S8) but not sulfate. 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. Among the anaerobic respirations, the reduction of sulfur species is most noteworthy because it gives rise to a toxic

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end product, hydrogen sulfide (H2S). Within the sulfur cycle, this end product serves as electron donor for a great diversity of aerobic chemotrophic and anoxygenic phototrophic microorganisms that may form visible blooms in sulfidic habitats. The sulfate- and sulfur-reducing prokaryotes do not form a phylogenetically coherent group, but members are found in several phyla within the domains Archaea and Bacteria. Energy generating mechanisms as well as other physiological, biochemical, and molecular biological aspects of this ancient group of organisms are described in detail.

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 substratelevel 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 nitrate reduction 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 are 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. This chapter gives an overview of prokaryotes that reduce sulfate or elemental sulfur in their energy metabolism (see > Fig. 9.1). Growth by reduction of other sulfur species is also included. Such bacteria have also been summarized as sulfidogenic bacteria (sulfide-forming) bacteria (Zeikus 1983; Lupton et al. 1984); however, strictly speaking, this term would also apply to putrefying bacteria that liberate sulfide from sulfurcontaining organic molecules during their degradation. Sulfate- or sulfur-reducing microorganisms are longestablished 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, this 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)

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Nitrogen compounds phosphate

Organic

Cell components

SO42– S0

Compounds (H)

CO2

Free H S 2 Energy

H2S

. Fig. 9.1 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

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 production resulted in the first isolation of a sulfate-reducing bacterium (named Spirillum desulfuricans), which was recognized as a strict anaerobe. The culture was isolated with malate and aspartate. Van Delden (1903a, b) grew sulfate-reducing 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 Stickland 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-50 -phosphosulfate (APS) was not

further phosphorylated to 30 -phosphoadenosine-50 phosphosulfate (PAPS), as in the assimilatory pathway, but rather directly reduced to sulfite and AMP (Peck 1959, 1962; Peck et al. 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 1954; Senez and LerouxGilleron 1954). 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 spore-forming strains were classified or reclassified in the new genus Desulfotomaculum (Campbell and Postgate 1965); the nonspore-forming, 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 35SO42 (Sorokin 1972). More than 50 % of the organic carbon in marine sediments was shown to be mineralized via sulfate reduction

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(Jørgensen and Fenchel 1974; Jørgensen 1977, 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 (Hoppe-Seyler 1886; Rubentschik 1928; Baars 1930), were confirmed to exist in sulfate-reducing 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, 1981b, 1982; Widdel 1980; Pfennig et al. 1981; 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, 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 CO-dehydrogenase 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. 1984, 1986, 1988; Powell et al. 1989) and genes for nitrogenase (Postgate et al. 1986; Kent et al. 1989), hydrogenase (Voordouw and Brenner 1985; Voordouw et al. 1985), 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 (Cypionka 1987, 1989; Warthmann and Cypionka 1990). Attempts to enrich acetate-oxidizing anaerobes with sulfur oxoanions 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 nutritional, morphological, chemical, or biochemical markers (Pfennig et al. 1981; 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 Pivovarova 1979). All other sulfate 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 d-subclass, 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 sulfatereducing 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 1988). An overview of the major groups of sulfate-reducing

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

bacteria and archaea (see following section) within the 16S rRNA-based tree of life is shown in > Fig. 9.2. 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 sulfate-reducing 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 (Roma˜o 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 cofactors, 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 1993). 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 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, 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-

9

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 sulfate-reducing 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 (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. Sulfatereduction 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 1999) and shown to constitute a significant fraction of the natural cold-adapted population (Sahm et al. 1999b). The combination of pureculture 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, sulfur-reducing, 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

313

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

a

Proteobacteria

1 Desulfctonaculum Desulfospoosinus

δ-Subclass Animals Chlaoflexus

Bacillus

3

Thermodesulfobacterium Thermotoga Thermodesulfovibrio Plants

Extreme halophiles

2 Archaeogbhus Methanogens

Sulfur-metabolizing and fermentative thermophiles

Microsporida

De robiu

acu

m ba

llis

arans

iab var ina

stre

vibrio d m esulfuri cans Desulfolomaculum nigrificans

Desulfolomaculum orientis

s

ob ulf

m

ii

ars

ba

rnu

infe

m

um

rvegicu s

dus no

lforhab

nus

ige

mn

ulu

arc

acin

lfo

su

ulfo

ii

lin

xo

ous a

b

Themro d osu

rha

a cle r

hicum utotrop rium a obacte Desulf Desulfobacter postgatel Desulfspira joergensenii Desulfobacula loluolica Desulfobulbus propionicus Desulfo capsa th iozymo Desulfo genes fustis g Desu lycolicu lforho Des s palus ulfb vacuo De acc latus aa su c eto lfo xida m ns on ile tie dje i

s

De

ulfo

ob

Des

Des

Sy ph

nlr o

Thermodesulfovibrio yellowstonli

o ap

otu

De

culatu

Thermodesulfotobacterium commune

s

ran

vo

lus

arc fos sul

sul

f

se

ml

Desud

en

onu

ns

fotr

lfomic

Desulfo

lba

ora

sul

Desu

re

ov

De

en

ium

og

lob

De

de

ha

s muitiv

hy

lfo

De

su

ococcu

rio

De

hal

vib

no

oph

tro

lla

na

lfo

su

b

lls oce

314

0.10 Stetteria hydeogenophila

Sulfolobus solfataricus Archaeoglobus fulgidus Methanobacterium formicicum

. Fig. 9.2 Phylogenetic trees reflecting the relationships of groups of sulfate-reducing bacteria to other organisms on the basis of 16S rRNA sequences. (a) Overview showing the three domains of life: (1) eubacteria, (2) archaebacteria, and (3) eukaryotes. The tree was adapted from Achenbach-Richter 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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 xcould be elucidated. It was recognized as an archaeal parallel of the CO-dehydrogenase pathway in mesophilic sulfate-reducing bacteria (Schauder et al. 1986, 1989), with the involvement of the archaeal cofactors (Mo¨ller-Zinkhan et al. 1989; Mo¨ller-Zinkhan and Thauer 1990; Schmitz et al. 1991; Klein et al. 1993; Schwo¨rer et al. 1993). Also, enzymes in the transport of reducing equivalents were investigated (Kunow et al. 1994, 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 sulfate-reducing 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 Tru¨per 1988; Dahl et al. 1990, 1993, 1994, 1999a; Speich et al. 1994; Sperling et al. 1998, 1999). In 1997, the complete genome sequence of A. fulgidus was published (Klenk et al. 1997). This was the first genome sequence of a sulfate-reducing 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 nineteenth 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 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 sulfur-sulfide cycle involving a green phototrophic sulfur bacterium that oxidized sulfide to

9

elemental sulfur and Desulfuromonas that reduced sulfur 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, 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. 1988). 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. 1995). A novel moderately thermophilic type of sulfur-reducing, acetate-oxidizing anaerobe was designated Desulfurella acetivorans (Bonch-Osmolovskaya et al. 1990). Furthermore, the thermophiles Aquifex (Huber et al. 1992), Ammonifex (Huber et al. 1996), and Desulfurobacterium (L’Haridon et al. 1995) 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 cataloging of Desulfuromonas (Fowler et al. 1986); it affiliates with the d-subclass 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

315

316

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

sulfur-reducing bacteria was termed the E-subclass of Proteobacteria. The first isolate (strain 5175; Wolfe and Pfennig 1977) was classified as Sulfurospirillum deleyianum (Schumacher et al. 1992). Desulfurella species were recognized as a distinct branch within the E-subclass with no specific relationship to sulfate-reducing bacteria or Desulfuromonas (Rainey et al. 1993; Miroshnichenko et al. 1998). The first biochemical studies of sulfur-reducing 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., Kro¨ger 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 Kro¨ger 1993; Schauder and Mu¨ller 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 (Schro¨der et al. 1988; Krafft et al. 1992, 1995; Schauder and Kro¨ger 1993). The three subunits of the polysulfide reductase were analyzed with respect to bound cofactors (e.g., molybdopterin, FeS centers) and the underlying genes (Krafft et al. 1992, 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, b; 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 Scho¨nheit and Scha¨fer 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. 1983). Further, newly isolated archaea that definitely grow by sulfur respiration, namely, on H2 and sulfur, were Stygiolobus 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, 1986) and Desulfurolobus (originally Sulfolobus) ambivalens (Zillig et al. 1985, 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 neutrophilus (Scha¨fer 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 nonphosphorylated 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 sulfur-respiring species Thermoproteus tenax and Pyrobaculum islandicum (Selig and Scho¨nheit 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 abyssi (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. 9.1. Hence, sulfate-reducing bacteria are easily recognized by the

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 amount 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 (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 papermaking 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, namely, 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 d-subclass 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. 9.2). 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 9.1.

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 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: 2CH3 CHOHCOO þ SO4 2 ! 2CH3 CHOO þ 2HCO3 þ HS þ Hþ DG 0 ¼

160 kJ=mol sulfate ð9:1Þ

2CH3 CHOHCOO þ 3SO4 2 ! 6HCO3 þ 3HS þ Hþ DG 0 ¼ 85 kJ=mol sulfate ð9:2Þ 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 sulfate reduction 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 1,323 kJ, respectively (here calculated 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. 9.1), as is obvious from relatively low growth yields. Examples of measured dissimilatory growth yields (YSulfate, cell dry mass formed per mol sulfate 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

317

Oval or rod

Oval

Oval or vibrio

Oval

Sphere

Oval (forms aggregates)

Rod

Multicellular filaments

Vibrio

Vibrio

Desulfomicrobium

Desulfobulbus

Desulfobacter

Desulfobacterium

Desulfococcus

Desulfosarcina

Desulfomonile

Desulfonema

Desulfobotulus

Desulfoarculus

Morphology

Vibrio

d

Desulfovibrio

Bacteria

Genus

Desulfoviridin 2

Optimum temperature ( C) +







35–39


34

30–32 

37

33

28–36 +

20–35


28–32


28–39


28–37


30–38 +

CO CO c c i

SO32
, S2O32
SO32
, S2O32
S2O32
, 3-Clbenzoate SO32
, S2O32
SO32
SO3 ,

CO

CO

S2O32


S2O32


CAC 

SO32
, S2O32


2


i

SO32
, S2O32
, NO3










+

+






+

+

i

SO32
, S2O32


+

H2

Acetate


+




Propionate








+ +

+

+





+



+

Ethanol




+

+

(+) (+) +




(+) +











f ND ND


(+) +

(+) +

(+) () 

+







Higher fatty acids

Electron donorsc

i

Electron acceptors for growth (other than SO4 2--) SO32 , S2O32 , fumarate

Oxidation of organic electron donorsb

. Table 9.1 Morphological and physiological properties of the genera of sulfate-reducing bacteria and archaea

Lactate


+






+






+

+









+












e


+

+

+

































Succinate, fumarate, and/or malate Fructose and/or glucose







+

+

+
























3- or 4-Anisate




Acetone

Methanol, glutarate, glutamate, phenol, aniline, nicotinate, indole










Methanol, glycerol, glycine, alanine, choline, furfural

Others utilized by some species

9

Phenyl-substituted organic acids

318 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Rod

Rod

Vibrio

Rod

Oval

Rod

Curved rod

Desulfohalobium

Desulfofustis

Desulfocella

Desulfocapsa

Desulfobacca

Desulfuromusa

Desulfospira

Vibrio

Vibrio

Desulfonatronovibrio

Desulfonatronum

Rod

Desulforhabdus

60

Oval

Oval

Desulfacinum

Desulforhopalus











c

i c

SO32
, S2O32


i

i

i

SO32
, S2O32
, S0 c

+







+










+




+

+












+




ND +










+

+

+ +

i




+

+

ND







i

S

+

ND

SO32
, SO32
, SO32
, SO32
, SO32
, SO32
, SO32
, S

+

c

SO32


0

+

i

S2O32
S2O32
S2O32
S2O32
S2O32
S2O32
S2O32
, 0

i

SO32
, S2O32


+

i

S2O32




i or CO

S2O32 Fumarate

ND S0, NO3
Fumarate c DMSO Fe(III)-citrate




26–30


30

37

20–30


34

28

37–40


37–40


37

37

18–19








60




Rod

65

Thermodesulforhabdus

Vibrio

Thermodesulfovibrio

30–37


65–70


Straight or curved rod, sporulates

Desulfosporosinus

30–38 50– 65h

Thermodesulfobacterium Rod

Straight or curved rod, sporulates

Desulfotomaculum




+










+










+

+

+















+

+

+










+

+















ND +

+

+

ND +

ND +

ND


ND +

ND +

+

+







ND +



+

+










+

+







+

+

+

+

+

+

+



+

+










+
















+

+







ND



ND




+


















+





















ND










+

ND

ND

ND










ND













2-methylbutyrate

Betaine, proline










L-Alanine,

Glycolate, betaine, choline, triethanolamine, indole
















Hexanol










3,4,5-Trimethoxybenzoate

Methanol, alanine

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

9 319

Rod

Rod

Desulfofaba

Desulfotalea

Archaeoglobus

Optimum temperature ( C)

Electron acceptors for growth (other than SO4 2--)








i i

SO32
, S2O32
SO32
, S2O32
Fe(III)-citrate CO

c

c

Oxidation of organic electron donorsb

SO32
, S2O32
Fe(III)-citrate

ND ND

Desulfoviridin 2

82–83


10

7

10

28

+

+









H2

Acetate


+













+




+

+

+

+

Ethanol +

+

+





g ND ND ND +







+

+

Lactate ND

+

+

+

+

+










ND










ND +

Starch, peptides










Toluene, p-cresol, benzaldehyde, benzoate, phenylacetate, p-hydroxybenzaldehyde, p-hydroxybenzoate

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, 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 et al. 1996), Desulfocella (Brandt et al. 1999), Desulfocapsa (Janssen et al. 1996), Desulfobacca (Oude Elferink et al. 1999), Desulfuromusa (Liesack and Finster 1994), Desulfospira (Finster et al. 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

Rod

Desulfofrigus

Sphere

Rod

Desulfobacula

Archaead

Morphology

Genus

Propionate

Electron donorsc

Higher fatty acids

. Table 9.1 (continued)

Succinate, fumarate, and/or malate Fructose and/or glucose

9

Phenyl-substituted organic acids

320 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

1981b); Desulfovibrio inopinatus, lactate (incompletely oxidized), 17.8 g (Reichenbecher and Schink 1997); Desulfococcus multivorans, benzoate (completely oxidized), 6.2 g (Widdel 1980); and 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 1996; Sass et al. 1998b; Knoblauch and Jørgensen 1999). 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); and 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 characterization of various species (for references see > Table 9.1). 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 ubiquitous 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

9

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 > Eq. 9.2.

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, 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 9.1). Other sulfate-reducing bacteria may produce some H2S in a by-reaction 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, b; 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 Desulfuromonas 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.

321

322

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 were 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. 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 9.2. 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-electron step process that occurs via a number of intermediates. However, unlike many nitrate-reducing bacteria, sulfate-reducing bacteria usually do not excrete the intermediate oxidation states, but only the final product sulfide. Only in two cases that 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 sulfate-reducing bacteria is driven by an ion gradient, as demonstrated in studies with Desulfovibrio species, Desulfobulbus propionicus, and Desulfococcus multivorans (Cypionka 1987, 1989, 1994, 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 103–104 (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 sulfate-reducing bacteria (Varma et al. 1983; Kreke and Cypionka 1992). With increasing sulfate concentration in the growth medium, the high-accumulating 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

Curved spiral

Sulfurospirillum deleyianum

25– 30

37

42 ND

i

i

Rod Rod Curved rods Rod Rod Rod Rod

Geobacter sulfurreducens

Pelobacter carbinolicus

Sulfurospirillum arcachonense

Hippea maritima

Desulfurobacterium thermolithotrophum

Aquifex pyrophilus

Ammonifex degensii

70

85

70

52– 54

26

35

35

35– 36

ND

+

+

+


f

+




+

+

+

+

+




ND

+

+

+




+

Formate

Succinate, fumarate, malate aspartate, oxaloacetate

Butyrate

Peptides, amino acids










Ethanol, propionate, succinate, glutamate

Others utilized by some species










+

+




+

+




ND




+




+

Formate

(With O2: sulfur, thiosulfate)

Ethanol, stearate, palmitate

Glutarate, glutamate

2,3-butandiol, acetoin, ethylene glycol

Ethanol, butyrate, succinate

ND ND Succinate, malate, glutamate
















+

NDb





+




Acetate +

Lactate and/or pyruvate

+







H2

c

i

i

ND

ND

Rod

Desulfitobacterium chlororespiransc

i

Rod

Vibrio

Dethiosulfovibrio peptidovorans

28– 37

i

c

Pseudomonas mendocina subsp.

Rod

Desulfomicrobium species

30– 36

52– 57

c

i

Vibrio

Desulfovibrio gigas

Optimum temperature (C) 30

Oxidation of organic electron donorsa

Wolinella succinogenes and similar spirilloid types Spirillum or vibrio 30– 37

Rod

Desulfurella acetivorans

Morphology Rod

a

Desulfuromonas acetoxidans

Bacteria

Species

d

Sulfur













+

+







+




























ND


+

+

+




+

+







+

+






























ND











+

ND +

+

+










+






Nitrate


+




+

ND ND ND ND +

+













+

+

+

+

+

+

+







Thiosulfate ND


+

+

+

+

+

+

+

+

+

+

+

+

+

+

+

Sulfite

Electron acceptors

Sulfate

Electron donors

Fumarate

. Table 9.2 Morphological and physiological properties of bacteria and archaea capable of respiratory reduction of elemental sulfur




+







+







+

+e

+



















O2

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

9 323

Long rod Disc with fibers Rod Lobed Coccus (irregular) 95 Disc

Pyrodictium occultum

Thermoproteus tenax

Stygiolobus azoricus

Stetteria hydrogenophila

Thermodiscus maritimus

+

+ + +


f f


f




+

+


f c

+

ND




+

f

Oxidation of organic electron donorsa
f

H2

Acetate




















Lactate and/or pyruvate


ND
















Yeast extract

Yeast extract




Yeast extract

(With O2: sulfur)

(With O2: sulfur)

Others utilized by some species

+

+

+

+

+

+

+

+

Sulfur

+




+







+




ND


+




+
















+

+




ND

ND ND


ND


ND


ND


ND


ND


ND




















O2

Symbols: c, complete oxidation (under anoxic conditions); i, incomplete oxidation; +, utilized; , utilized or not utilized;
, not utilized; ND, not determined or not reported a 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 et al. 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 Scho¨nheit 1994); Pyrodictium occultum (Fischer et al. 1983; Stetter et al. 1983); Thermoproteus tenax (Zillig et al. 1981; Fischer et al. 1983; Scha¨fer et al. 1986; Stetter et al. 1990; Selig and Scho¨nheit 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 fermentative metabolism e Low partial pressure (microaerobic conditions) f Obligate lithoautotrophs that do not oxidize organic compounds

85

80

80

105

100

88

Lobed coccus

Pyrobaculum islandicum

90

Sulfolobus ambivalens

Morphology Lobed coccus

a

Optimum temperature (C)

Acidianus infernus

Archaea

Species

Thiosulfate

Electron acceptors

Sulfite

Electron donors

Sulfate

. Table 9.2 (continued)

Fumarate

9 Nitrate

324 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

to attenuate even the low-accumulating system to prevent excess buildup of sulfate. Most likely, the sulfate-transport systems operate near equilibrium (Cypionka 1994, 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). Sulfate 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 (E00 = 0.516 V) than redox potentials of most catabolic redox couples (> Fig. 9.3). Before being reduced, sulfate is activated by ATP sulfurylase (Peck 1959, 1962); the product is adenosine-50 -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 30 -phosphoadenosine-50 -phosphosulfate (PAPS) before reduction (Trudinger and Loughlin 1981; Fischer 1988; Peck and Lissolo 1988). 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: SO4 2 þ ATP þ 2Hþ ! APS þ PPi DG 0 ¼ þ46 kJ=mol

ð9:3Þ

PPi þ H2 O ! 2Pi DG 0 ¼
22 kJ=mol

9 ð9:4Þ

Sum reaction: SO4 2
þ ATP þ 2Hþ þ H2 O ! APS þ 2Pi DG 0 ¼ þ24 kJ=mol

ð9:5Þ

High pyrophosphatase activities were found in Desulfovibrio (Fauque et al. 1991), Desulfobulbus (Kremer and Hansen 1988), and 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 1989). 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 > Eq. 9.5. 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 channeled 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. 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; Kukko-Kalske et al. 1989) to allow formation of a substantial APS concentration. However, ATP sulfurylase 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, 1999). The ATP sulfurylase in E. coli, therefore, has been termed the ‘‘ATP sulfurylaseGTPase system.’’ The assimilatory ATP sulfurylase from E. coli and the dissimilatory enzyme from Desulfovibrio species also

325

326

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

E (V)

E (V)

–0.5

–0.5

–0.4

–0.4

NADP+

–0.3

–0.3

Lactate

Pyruvate

–0.2

–0.2

Fdll(red)

Fdll(ox)

–0.1

–0.1

Pyruvate

AeCoA + CO2

Fd(red)

Fd(ox)

H2 (10Pa)

H+

H2 (10Pa)

H+

NADPH

MKH2 Succinate

MK

SO42–

SO3–

SO3–

H2S (6e)

Electrons Fumarate

–0.0

–0.0

+0.1

+0.1

+0.2

+0.2

+0.3

+0.3

(2e) APS

HSO + 3–AMP

. Fig. 9.3 Comparison of redox potentials of some important electron-donating and electron-accepting reactions in sulfate-reducing bacteria. As mechanism of sulfite reduction to sulfide, a direct reduction with six electrons (6e
) is assumed. For ferredoxin, an average of the E00 (
0.440 V) given by Fauque et al. (1991) and the E00 (
0.400 V) given by Thauer (1988) and Thauer et al. (1989) is indicated. Abbreviations: Fd ferredoxin, MK menaquinone

differ markedly on the structural level. A recent study on the composition of ATP sulfurylase from two sulfate-reducing 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 E0 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 Tru¨per 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 Tru¨per 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; Tru¨per 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 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 1982; > Fig. 9.4). 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 (Mu¨ller 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

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes NH2 N

N

O− O− 5′ − H2C O P O S O

N

N

OH

OH

O

O

O H H3C

Adenosine H+

APS

O

O− O P

−

O

O

O S

O

O

N

H3C

N

O−

NH

5

NH

1 N

H 3C

O

N H

H3C

N

R

O

N H

R

FAD-reduced

NH2

2 [4Fe-4S]2+

N

N 2 [H]

O 5′ H2C O P

N

N

OH +

+

2 [4Fe-4S] + 2H

O O −

O

S

O H3C

N

N R

N

AMP O O

H3C

NH

H 3C

OH

OH

O

N NH

O

H 3C HSO3

–

+

H

FAD oxidized

N

N H

O

R FAD-sulfite adduct

. Fig. 9.4 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 FAD-sulfite 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-50 -phosphosulfate, FAD flavin adenine dinucleotide

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. 9.5). 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 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 bisulfites (tautomeric forms, [:SO2O-H] and [H-SO2O:] ), which are approximately equally abundant at pH 7.0 (pKa2 = 6.99), are pyramidal

327

328

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

O

O Adenosine

O P O O−

O Adenosine

−O

S S

Enzyme

O−

O

O P O O−

O

S O−

Adenosine

O P

O−

O−

O

Aps

AMP

O −S

−O

Enzyme SO32−

S S

Enzyme

O−

2e−

. Fig. 9.5 Proposed mechanism of adenosine-50 -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 (2e ) allows separation of sulfite and regeneration of free thiolate

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 (> Eq. 9.6). SO3 2 þ 6e þ 8Hþ ! H2 S þ 6H2 O

S

S

S

H2C

Fe

Fe

CH2

S Fe S

S Fe

S

H2C HO2C

CO2H S

ð9:6Þ

The active centers of dissimilatory and assimilatory sulfite reductases (and nitrite reductases) are characterized by two metallo-cofactors, a reduced porphyrin of the isobacteriochlorin class, the siroheme (Murphy and Siegel 1973; Murphy et al. 1973b, 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. 9.6. Siroheme-containing reductases have been isolated from a wide range of organisms. Siroheme was identified in assimilatory sulfite reductase from Escherichia coli (Murphy et al. 1973a), dissimilatory sulfite reductase from Desulfovibrio species (Murphy et al. 1973b), the dissimilatory ‘‘reverse’’ sulfite reductase of thiobacilli (Schedel et al. 1975; Tru¨per 1994) and Chromatium (Schedel et al. 1979), and the ammonium-producing 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 Garrett 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

HO2C

CO2H N

N Fe

H

N

CH3

N

CO2H HO2C CH3

H

CO2H HO2C

SO32–

. Fig. 9.6 Prosthetic group of sulfite reductase. Two metallo-cofactors, 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 nonbridging side

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

desulforubidin and desulfofuscidin, and P582 (> Table 9.3; 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 g-subunit is not encoded in the same operon as the a- and b-subunits and is not expressed coordinately with the a- and b-subunits (Karkhoff-Schweizer et al. 1993). The molecular mass of 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 sirohydrochlorin 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, 1959). Siroheme prepared from desulfoviridin was found to catalyze the reduction of sulfite to sulfide and thiosulfate in the presence of artificial 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 Mo¨ssbauer 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, c). 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 supercluster (Fritz 1999). Based on these findings, it is supposed that resonance signals previously thought to originate from a supercluster 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

9

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 variabilis (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 sulfate-reducing 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 (desulfoviridin 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. 9.7): a sequential reduction in three twoelectron 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. 9.7b), is mostly based on in vitro studies (Kobayashi et al. 1972; Akagi 1983). In the in vitro experiments, methyl- or benzylviologen was 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, namely, 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 ‘‘thiosulfate-forming’’ 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):  2 þ 2e H35 SO3 þ O3 S35 S SO3  35 35 2 ! O3 S S þ HSO3 þ SO3 2

ð9:7Þ

Also purified from Desulfovibrio vulgaris was a ‘‘trithionatereducing 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

329

19(3) [Fe4S4](2) [Fe6S6](3)

10(4), 22(3)

10(4), 18(3)

[Fe4S4](4) [Fe6S6](5)

Total iron

Acid-labile sulfur

[FeS] clusters

Pierik et al. (1992a)

Steuber et al. (1995)

Wolfe et al. (1994)

(2)

(3)

(4)

Akagi (1983)

Lee and Peck (1971)

(6)

(7)

Pierik et al. (1992b, c)

(5)

Lee et al. (1973a)

(1)

References

Lee et al. (1973b)

Arendsen et al. (1993)

(3)

Moura et al. (1988)

(2)

(1)

Methylviologen Hatchikian and Zeikus (1983)

(1)

(1)

Methyl- or benzylviologen

Methylviologen

(1)

S3O62 , NH4+(1,2) S2 , S2O32

(1)

+ SO32 , NO2 NH2OH(2)

(1)

Electron donor (in vitro)

(1,3)

S2 , S3O62

SO32

(1)

[Fe4S4]

16–17(1)

21(1), 32(2)

4(1,2)

167(1) 190(2)

a2b2(1)

720, 580, 545, 392(1) 693, 576, 389, 279(1)

+

P582

alSiRa

(1,2)

Akagi and Adams (1973)

(2)

Trudinger (1970)

(1)

(1)

(2)

Methylviologen

S3O62 , S2O32

S2

SO32 , NO2 NH2OH(1)

+

(1)

15 matoms per g protein

54 matoms per g protein

145(1)

700, 582, 392, 280(1)

(3) (3)

(1,2)

S2O32 , S2O3

S2

SO32 , NO2

[Fe4S4](1)

20(1), 47(3)

24(1), 51(3)

160(1) 280(3)

a2b2(1)a4b4(3)

700(3), 594, 393, 274(1)

Thiobacillus Chromatium

Reverse SiR

Dahl et al. (1994)

(2)

Schedel et al. (1979)

(3)

Schedel et al. (1975)

(2)

Triper (1994)

(1)

Methylviologen Methyl- or benzylviologen

SO32

[Fe4S4]

20(2)

22–24(1)

2(1)

178.2(1)

a2b2(1)

593, 545, 394, 281(1)

Archaeoglobus

Archaeal SiR

Moura and Lino (1)Dahl et al. (1994) (1993)

(1)

Methylviologen

S2

SO32

+

[Fe4S4](1)

5(1)

5(1)

1(1)

27(2)

Monomer

590, 545, 400(1)

Thermodesulfobacterium Desulfotomaculum Desulfovibrio

Desulfofuscidin

(6,7)

S3O62 , S2O32

Minor products

(1)

S2

Major products

, NH4+(4)

Known substrate(s) SO32 (1), NO2 NH2OH(4)

aSir, sulfite reductase

a

15(3), 21(2)

2(4)

Number of sirohemes

(4)

2(2)

226(1)

Molecular weight (kDa)

Reaction with CO

225(1)

a2b2 (%)(2,3) (n = 1 3)

Subunit structure

(1)

a2b2a2b2 (%)(2,3)

628, 580, 408, 390, 279(1)

Absorption maxima (nm)

Desulfovibrio Desulfosarcina

Desulfovibrio

Organism

Desulforubidin

Desulfoviridin

Properties

Siegel et al. (1982)

(2)

Siegel and Davis (1974)

(1)

NADPH methylviologen

S2 , NH4+

SO32 , NO2 NH2OH(1)

[Fe4S4]

4(1/b)

685

a2b2(1)

714, 587, 386, 280(1)

Escherichia

aSiR

9

Type of sulfite reductase

. Table 9.3 Biochemical characteristics of sulfite reductases

330 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

a

6 e−

SO32−

3 H2O H2S

8 H+

b

2 e− 3 H2O

3 SO32−

2 e−

6 H+

2 e− S2O32−

S3O62− SO32−

9

H2 S 2 H+ SO32−

. Fig. 9.7 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

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. 9.7b). 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 (E00 = +0.225 V) and thus a favorable acceptor even for high-potential electron donors, as, for instance, from dehydrogenation of succinate (E00 = +0.030 V). The nature of the natural electron donor of the three two-electron reduction steps of the trithionate pathway has not 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 oxoanions. Trithionate and thiosulfate reductases may serve for utilization of their substrates from the environment or for scavenging them as by-products of the bisulfitereductase reaction. Low concentrations (5–100 mM) of thiosulfate formed in de-energized cells from added sulfite (Fitz and Cypionka 1990) or in cells growing on sulfate (Vainshtein et al. 1980) also may be interpreted as by-products resulting from a reversely operating thiosulfate reductase with sulfide and sulfite as reactants; the electrons from this low-potential reaction (E00 = 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 sulfurcontaining 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 sulfatereducing 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, b; Moura and Lino 1994). Tan and Cowan (1991) proposed a mechanism for 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 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. 9.8). 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

331

332

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes Fe2+ S

Fe3+

S

S

Fe2+

Fe3+

SO32–

Fe2+ H+

L

Fe2+

HO

O−

S

OH−

HO

Fe2+ 2 e−

S Fe2+

O−

S ..

O

S ..

OH

O

H2S OH−

2 H+

Fe2+

Fe3+

Fe2+

S

S

Fe2+

Fe3+

Fe2+

:S: ..

:S: ..

S

2 e−

OH–

:S: OH

Fe3+ 2 e−

S Fe3+

H+

:S: O

. Fig. 9.8 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))

siroheme into the sulfite. In addition, the local environment in the sulfite-binding 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 hemecontaining oxygenases (Dawson 1988; Poulos 1988). Lui and Cowan (1994) have also proposed a six-electron reduction via a push-and-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 has 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, 1982; Siegel and Davis 1974). 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 A˚ 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 A˚ (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 A˚ structure also allowed recognition of further refined details of the structure. The hemoprotein consists of three 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. Domains 1 and 10 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 protonrich 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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 (domains 10 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 NO-forming nitrite reductase in denitrifiers), which catalyzes the dissimilatory reduction of nitrite to ammonia (> Eq. 9.8). Such types of nitrite reductases also contain siroheme in the active center. þ

þ

NO2 þ 6e þ 8H ! NH4 þ 2H2 O

ð9:8Þ

Cytochrome c nitrite reductase from Sulfurospirillum deleyianum not only catalyzes the six-electron 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 ten closely arranged hemes. Apparently, different structures and probably also varied mechanisms have evolved to accomplish a six-electron reduction. A membrane-bound cytochrome c-containing nitrite reductase (also isolated from Desulfovibrio desulfuricans) catalyzes the six-electron 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, Archaeoglobus 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

9

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). 4SO3 2 þ Hþ ! 3SO4 2 þ HS DG 0 ¼
58:9 kJ=mol sulfite

ð9:9Þ

S2 O3 2
þ H2 O ! SO4 2
þ HS
þ Hþ DG 0 ¼
21:9 kJ=mol thiosulfate

ð9:10Þ

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 þ 4H2 O ! SO4 2
þ 3HS
þ 5Hþ

ð9:11Þ

Standard concentrations: pH = 7: DGo0 = +10.2 kJ/mol sulfur SO42
and HS
, 0.001 M; pH = 7: DGo0 = +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 (> 9.9) and (> 9.10) 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 (Kra¨mer 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 1988,

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 (E00 = 0.060 V; see also > Fig. 9.3). 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 (Kra¨mer 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 9.1). 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 section > ‘‘Sulfur-Reducing 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, 1980; Cammack et al. 1984). Sulfonates, DMSO

Reduction of sulfonates by sulfate-reducing bacteria was first described by Lie et al. (1996). These authors demonstrated utilization of cysteate, isethionate (2-hydroxy-ethanesulfonate), 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 sulfate-reducing 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 74-kDa protein with a [4Fe-4S] center and a molybdopterin 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 nonsulfate-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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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. 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 suggests that permanently oxic waters or soil usually does not harbor nonspore-forming 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, nonspore-forming sulfate-reducing 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 sulfate-reducing 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

9

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 sulfate-reducing 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 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– 0.48 mM was 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 (Marschall 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, 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).

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 sulfate-reducing bacteria comes from the observation of aerotaxis in Desulfovibrio species. In medium 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 sulfatereducing 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 methyl-accepting 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 C-terminal 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 knockout 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 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 nonheme iron proteins that have been purified from Desulfovibrio species found to catalyze removal of superoxide (Roma˜o 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 above-mentioned 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 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; Kro¨ger 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.

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 tiedjei (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 2,4,6-tribromophenol to phenol (Boyle et al. 1999); other halogenated compounds that are used as alternative electron acceptors include 2-, 4-, 2,4-, and 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 sulfate-reducing 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 (1999). 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 terminal-oxidase 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 sulfatereducing 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

9

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. 1988b). 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 redoxcarriers 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 section > ‘‘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). 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 cellfree 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 A˚. The three cytochromes had a similar overall

337

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 IVare 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 toward either [Fe] or [NiFe] hydrogenase (Pieulle et al. 1996; Magro et al. 1997). An octaheme cytochrome c3 that is found in most Desulfovibrio species is structurally rather different from tetraheme cytochrome c3 but also can react with hydrogenase (Fauque et al. 1991; 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, 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 domains 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 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 nonheme iron-containing 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; Blanchard et al. 1994; Sebban-Kreuzer et al. 1998a, b). 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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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).

9

1988; Brumlik and Voordouw 1989). Kremer et al. (1988a) 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 section > ‘‘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 section > ‘‘Oxygen’’ in this chapter).

Ferredoxins

Ferredoxins are very common in sulfate-reducing and sulfurreducing 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 (E00 = 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 appropriate electron donor for the thiosulfate reductase reaction. Ferredoxin II (E00 = 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. 1989). Flavodoxins

In some but not all Desulfovibrio and Desulfomicrobium species, flavodoxins have been found. The two oxidation states, F/FH (E00 = 0.140 V) and FH/FH2 (E00 = 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 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 are 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, b; 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

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 (E00 = +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 (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 (Kro¨ger et al. 1988; Mo¨ller-Zinkhan and Thauer 1988; Mo¨ller-Zinkhan and Thauer 1989; Thauer et al. 1989; > Figs. 9.6, > 9.8) 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 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).

339

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 sacrificed for the disposal of surplus reducing equivalents (e.g., regeneration of NADP+). With external electron acceptors, substrate-level phosphorylation and growth are 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 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 sulfate-reducing bacteria, numerous enzymes catalyzing redox reactions as well as potentially electron-carrying 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 sulfate-reducing 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 9.1) reflects the ecological importance of the lightest of all molecules. Hydrogen at standard pressure is an energetically favorable electron donor (2 H+/H2, E00 = 0.414 V); the free energy change at various partial pressures is depicted in > Fig. 9.11. Cell material during growth on H2 and sulfate may be synthesized from acetate and CO2 (chemolithoheterotrophic species) or alone from CO2 (see autotrophic species; section > ‘‘Carbon Assimilation’’). Growth on H2 has been observed in many of the known genera of sulfate-reducing bacteria (> Table 9.1). Hydrogenase

activities have been demonstrated in strains of the genera Desulfovibrio (Fauque et al. 1991), Desulfobulbus (Samain et al. 1986b; Kremer and Hansen 1988), 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 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 cocultures (see section > ‘‘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: H2 Ð Hþ þ H Ð 2Hþ þ 2e

ð9:12Þ

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 9.4) 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 was 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 9.8). 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 sulfate-reducing bacteria were

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

. Table 9.4 Brief overview of characteristics of different types of hydrogenases found in Desulfovibrio species [Fe] [NiFe] hydrogenase hydrogenase

[NiFeSe] hydrogenase

Catalytic activity H2 uptake

Very high

Moderate

Low

H2 evolution

High

Moderate

Moderate

CO

Very high

High

Moderate

NO

Very high

High

Very high

Sensitivity to

Nitrite

Moderate

No

Moderate

Acetylene

No

High

Moderate

90

81

Molecular mass 57 (kDa) Adapted from Fauque et al. (1991)

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 [4Fe-4S] clusters were identified. The atypical Fe cluster, 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 three-dimensional structure revealed that this hydrogenase displays a novel protein fold and that the H cluster is composed of a typical [4Fe-4S] cluster 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 resides 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

9

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 sulfatereducing 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 H2-activating 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 (De Luca 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 (De Luca et al. 1998b). Thus, the NADP+-reducing hydrogenase appears to differ structurally from the three other types of hydrogenases. The [NiFe] hydrogenases have been purified from D. desulfuricans (Kru¨ger et al. 1982), D. gigas (Moura et al. 1982), D. multispirans (Czechowski et al. 1984), and D. africanus (Nivie`re 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 A˚ resolution (Volbeda et al. 1995, 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 A˚. 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

341

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

O N

N C

C

C

Fe Cys

S

S

Cys

Ni S Cys

. Fig. 9.9 Suggested prosthetic group of [NiFe] hydrogenases. Three dinuclear, nonprotein ligands (2 CN , 1 CO) coordinate the Fe atom in the active center (Happe et al. 1997; Higuchi et al. 1997; Pierik et al. 1999)

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 A˚) 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 CN as dinuclear ligands of the NiFe 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. 9.9; Happe et al. 1997); the function of the dinuclear ligands remains unclear. The three [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 sulfate-reducing 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] 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. 1988). 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 selenocysteyl-tRNASec is synthesized from seryl-tRNASec and selenophosphate by selenocysteine synthase (Bo¨ck et al. 1991; Heider and Bo¨ck 1993). SelenocysteyltRNASec 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 selenocysteine-inserting 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 Bo¨ck 1997). First evidence for a periplasmic location of [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 is often localized in the periplasm. Investigated species are Desulfovibrio vulgaris strain Marburg (Badziong and Thauer 1980), Desulfovibrio vulgaris

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

strain Hildenborough (van der Westen et al. 1978), Desulfovibrio desulfuricans (Steenkamp and Peck 1981), Desulfovibrio gigas (Bell et al. 1974), and 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 1986; 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 periplasmorientated 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 b-lactamase from Escherichia coli lacking its own leader sequence allowed export of the enzyme. Exchange of one of the two arginines in the leader sequence to glutamate by sitedirected mutagenesis inhibited export of b-lactamase completely (Nivie`re 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

9

(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 are 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 alteration of the hydrogenase-cytochrome interaction (Aubert et al. 1997, 1998a). The positive charges surrounding the surface-exposed 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 [FeS] clustercontaining 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 wild type 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). Similarly, the [NiFe] hydrogenase from Desulfovibrio desulfuricans can reduce the

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 hydrogenase 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 13/4–21/3 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. 9.10). Because sulfate activation is associated with a net consumption of 2 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 3 H+/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 sulfatereducing bacteria). Nothing is known about the possibility of a Q-cycle (Peck and Lissolo 1988); considering this translocates

+ + +

– – –

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/3 ATP

H2S

x H+ H2S + + +

– – –

. Fig. 9.10 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-50 -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

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 sulfate-reducing bacteria with H2 over a wide range of partial pressures. At standard pressure, H2 is one of the energetically most favorable electron donors (DG0 =
152.2 kJ/mol sulfate). However, Desulfovibrio was shown to scavenge H2 below 10 Pa (10
4 atm.; Cord-Ruwisch et al. 1988).

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

donor that is equivalent to H2. The redox potential of the couples 2 H+/H2 and HCO3 /HCOO is very similar (E00 around 0.41 V). Hence, formate is a favorable electron donor:

G (kJ / mol CH or SO)42–

50 0 –50 –100

4HCO2 þ SO4 2 þ Hþ ! 4HCO3 þ HS DG 0 ¼
146:6 kJ=mol sulfate

SO42– reduction H2 formation from CH4

with H2

–150 –200 10 –3 10 –2 10 –1

10 8

9

10 7

10 6

1

10 1 10 2 (Pa)

10 3

10 4

10 5

10 5

10 4 10 3 (An)

10 2

10 1

1

H2 partial pressure

. Fig. 9.11 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

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. 9.11. Assuming that net ATP synthesis coupled to any catabolic overall 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. 1988; Thiele and Zeikus 1988); formate transfer was most likely to occur in a sulfatereducing 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

ð9:13Þ

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 [FeS] center is similar to the [FeS]-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. 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 membrane. 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.

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Terminal Oxidation and Utilization of Acetate

The oxidation of organic substrates in sulfate-reducing 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, namely, the presence of a pathway for acetyl-CoA oxidation, usually includes also the ability to use free acetate as a growth substrate: CH3 COO þ SO4 2 ! 2HCO3 þ HS DG 0 ¼
47:6 kJ=mol sulfate

ð9:14Þ

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. 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, n-butanol, or isobutanol to propionate, butyrate, or isobutyrate, respectively (Mechalas and Rittenberg 1960), and the formation of propionate from C-odd fatty acids (Pfennig and Widdel 1981; 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 14 C-labeled substrates. In Desulfobacter postgatei, all enzymes of a citric-acid cycle were found (Brandis-Heep et al. 1983; for an overview, see Kro¨ger et al. 1988; Thauer 1988, 1989; Thauer et al. 1989). Also with position-labeled [14C]-acetate as growth substrate, the labeling pattern in aspartate and glutamate that are derived from oxaloacetate and 2-oxoglutarate was 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. 9.12). 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 acetylCoA and oxaloacetate to citrate in Desulfobacter is associated with ATP synthesis (Mo¨ller 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 ATPcitrate lyase reaction is reversible (DG0 = 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 malate dehydrogenase is neither NAD+ nor NADP+. The reduction of the artificial acceptor 2,6-dichlorophenol indophenol (DCPIP) was inhibited by the menaquinone antagonist 2(n-heptyl)-4hydroxyquinoline-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 (Mo¨ller-Zinkhan and Thauer 1988). Still, it is most likely that Desulfobacter employs a more positive acceptor for malate oxidation than other bacteria. The reversible, energyconserving 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. 9.12. 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

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes Acelate ATP ADP + P1

a

b

ADP ATP + P1 X[H]2

AcCoA MADH

Citr

OxAc

MAD(P)H OxAc

Citr

OxAc

+

Icitr

AcCoA

H2O Fum MK

Succ

Acelate

CO2

MAD(P) Icitr

Mal

Mal

MADP+

MADP+

MADPH CO2

MADPH

Z-OCl Fd(ox) SucCoA Fd(red)

AcCoA

H2O

CO2 Fum

MKH2

Citr

+

MAD

X Mal

MKH2

c

MK

2-OGl Fd(ox) SucCoA Fd(red)

Succ

Acelate

Icitr MADP+ MADPH CO2

H2O Fum X[H]2

X

Succ

2-OGl Fd(ox) SucCoA Fd(red)

CO2

CO2 ATP ADP +P1

. Fig. 9.12 Modifications of the citric-acid cycle for the anaerobic oxidation of acetate in three species of sulfate- and sulfur-reducing bacteria: (a) Desulfobacter postgatei, (b) Desulfuromonas acetoxidans, and (c) Desulfurella 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. (1989))

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, namely, 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. 9.13). 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+ was the 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, tetrahydropteroyltetraglutamate, which has four glutamate residues instead of one (La¨nge 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 (Mo¨ller-Zinkhan et al. 1989). The bioenergetic implications of the pathways for acetate oxidation and terminal oxidation of other organic compounds

in sulfate- and sulfur-reducing bacteria have been discussed (Kro¨ger et al. 1988; Thauer 1988; Thauer et al. 1989). The free energy gain from reduction with acetate (> Eq. 9.14) is much lower than from sulfate reduction with H2 at standard pressure (> Fig. 9.11). 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 = /) 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 h. 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 (> 9.14) with the generally observed requirement of >70 kJ/mol ATP (Schink 1988a; Thauer et al. 1977) again indicates that acetate-oxidizing 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;

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Organic electron donor

a

Organic electron donor

b

O

O — —

— —

—C— — SCoA CH3 —

—C— — SCoA CH3 —

CH3 — THP

[CO]

[CO]

CH3 — THMP

X

F420

Quinonered

X[H]2

F420H2

Quinoneox

— THMP CH2 —

— THP CH2 — NAD(P)+ NAD(P)H — THP CH —

X

F420

Quinonered

F420H2

Quinoneox

CH — — THMP

X[H]2

X X[H]2

CHO — THMP

CHO — THP ADP + Pi ATP

CHO — MF

HCOO−

X

NAD+

X[H]2

NADH CO2

CO2

CO2

CO2

. Fig. 9.13 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. (1989))

however, 2 ATP are needed for sulfate activation (assuming that pyrophosphate is irreversibly hydrolyzed). The energy requirement for sulfate transport in 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 (DE00 = 0.25 V) could be associated with a translocation of 2 H+/2 [H], which are 4 H+/acetate (Kro¨ger et al. 1988; Thauer 1988). The preceding

reduction of NADP+ with ferredoxin (DE00 = 0.1 V) has been discussed as another energy-conserving step allowing the translocation of 1 H+ (> Fig. 9.14a). 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 (E00 = +0.033 V) with menaquinone (E00 = 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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

a

b

E′ (V)

–0.4

c

2-OGl

2-OGl

Fd(red)

Fd(red)

Icitr

~ +? Δµ H

9

[CO]

HCOO– ~ +? Δµ H

Icitr NADH

NADPH

–0.3

~ +? Δµ H

NAD NADPH

CH2=THF ~ +? Δµ H

~ +? Δµ H

–0.2 Mal ?

Y Mal

[S]

X

HSO3–

HSO3–

CH3=THF

–0.1 MKH2

MKH2

MKH2

APS

0.0 Succ

APS

Succ

. Fig. 9.14 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 E00 ); 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 HSO3 and a range of 1–10 mM H2S (dotted redox span). The concentration range of H2S for sulfur reduction is also 1–10 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-50 -phosphosulfate. Fd(red), reduced ferredoxin; Icitr, isocitrate; Mal, malate; Succ, succinate; THF, tetrahydrofolate; 2-OGl, 2-oxoglutarate (Adapted from Mo¨ller-Zinkhan and Thauer (1988) and Thauer (1988))

a mere thermodynamic viewpoint, any unfavorable partial reaction is rendered possible if embedded in an exergonic overall reaction. In biological systems, 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 primary 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 gradient-driven 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 membrane-bound 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 (Kro¨ger et al. 1988; Thauer 1988). With the remaining one to two protons, the net energy conservation in the sulfur reducer would be 1 /3 to 2/3 ATP per molecule of acetate. Suggested proton-translocating reactions in the C1/COdehydrogenase pathway are indicated in > Fig. 9.14c. Propionate

Propionate serves as electron donor and carbon source for the incompletely oxidizing Desulfobulbus species and several completely oxidizing sulfate reducers (> Table 9.2). Propionate in Desulfobulbus is oxidized to acetate via a randomizing

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Ethanol

Lactate

Propionate ATP ADP Propionyl- P 2X

Succinyl-CoA

2X[H]2

2X

ADP + Pi ATP

Propionyl-CoA 2X[H]2 Methylmalonyl-CoA TC COO–

TC

Succinate 2X

• •

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. 9.15 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

pathway with succinate, a symmetric molecule, as free intermediate (Kremer and Hansen 1988; > Fig. 9.15). 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 section > ‘‘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 nonheme 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/COdehydrogenase 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

Butyrate and higher fatty acids are oxidized by many incompletely and completely oxidizing sulfate-reducing bacteria (Widdel 1980; Pfennig and Widdel 1981; Widdel 1988; > Table 9.2). 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: n n CH3 ðCH2 Þ2n COO þ SO4 2 ! ðn þ 1ÞCH3 COO þ H2 S 2 2 ð9:15Þ n CH3 ðCH2 Þ2n þ 1 COO þ SO4 2 ! nCH3 COO 2 n þ CH3 CH2 COO þ H2 S 2

ð9:16Þ

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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

group; this leads to formation of propionyl-CoA rather than acetyl-CoA 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: 4HðCH2 Þn COO þ ð3n þ 1ÞSO4 2 þ ð2n þ 2ÞHþ ! ð4n þ 4ÞHCO3 ð3n þ 1ÞH2 S

ð9:17Þ

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 succinyl-CoA 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 sulfatereducing bacteria, is utilized by most species of almost each genus and may be oxidized completely or incompletely (> Eqs. 9.1 and > 9.2, 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 membrane bound. None of the enzymes from sulfatereducing bacteria have been purified to homogeneity. D-Lactate

9

dehydrogenase of Desulfovibrio desulfuricans was present in the particulate fraction, and detergents were required for its solubilization (Czechowski and Rossmore 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 1982), 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 oxidoreductase (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, b; 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, acetylCoA 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 sulfate-reducing 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 (> Eq. 9.1) 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 section > ‘‘Activation of

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 so-called hydrogen-cycling model for growth on lactate of a Desulfovibrio involved the cytoplasmic production of H2 as a result of the oxidation of 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 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 H2 transfer 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 H2-evolving 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 9.1). 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 sulfatereducing bacteria form strong smelling by-products 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 lactategrown 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, 1995a). The first 21 N-terminal 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 so-called 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 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. 1988a). 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. 1988a). Furthermore, it was shown that acetaldehyde can be oxidized in Desulfovibrio gigas by two completely different enzymes, a molybdenumcontaining 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. 1988a). The synthesis of the enzymes appeared to be strongly affected by the presence of molybdate and tungstate in the growth media

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

(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 benzylviologen-linked tungstencontaining 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 molybdenumcontaining aldehyde oxidoreductase is a homodimer of subunits with 907 amino acid residues and contains a molybdopterin cofactor and two different [2Fe-2S] 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; Roma˜o et al. 1995). The tungstencontaining aldehyde oxidoreductase of Desulfovibrio gigas was active toward 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; Roma˜o 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-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 2-propanol 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

9

a 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 3-hydroxypropionate or to acetate production as the major product (Nanninga and Gottschal 1987; Qatibi et al. 1991; Tanaka 1990, 1992); oxidation of 1,4-butanediol and 1,5-pentanediol yielded the corresponding hydroxyacids (Tanaka 1992). Oppenberg and Schink (1990) suggested a pathway involving malonylsemialdehyde for the conversion of 1,3-propanediol 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 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 biacutus. 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

353

354

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 sulfate-reducing bacteria was described that could utilize the long-chain ketones hexadecan-2-one and 6,10,14-trimethylpentadecan-2-one. The oxidation of these ketones also involved a carboxylation reaction (Hirschler et al. 1998). Glycolate

Glycolate is a widespread by-product 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).

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 (Imhoff-Stuckle 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 Desulfobacterium 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).

Malate, Fumarate, Succinate, and Other Dicarboxylic Acids

Furfural

Dicarboxylic acids known from the citric-acid cycle are relatively common substrates of incompletely and completely oxidizing sulfate reducers (Postgate 1984a, b; 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, E00 = +0.033 V). In various Desulfovibrio strains, the C4-dicarboxylic 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).

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 13C-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.

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 and

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 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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 sulfatereducing 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 sulfate-reducing 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, b). Several new types of sulfate-reducing 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 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 9.5. 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 sulfatereducing bacteria are therefore expected to be completely different from those of aerobic bacteria and to involve novel biochemical reactions. Most insights into the degradative

9

. Table 9.5 Sulfate-reducing bacteria with the capacity to use aromatic compounds as growth substrates Organism

Aromatic substratea

Reference

Non-hydrocarbon aromatic compounds Desulfonema magnum

Benzoate, 4-hydroxybenzoate, phenylacetate, 3-phenylpropionate, hippurate

Widdel et al. (1983)

Desulfococcus niacinib

Nicotinic acid, 3-phenylpropionate

Imhoff-Stuckle and Pfennig (1983)

Desulfobacterium indolicumc

Indole, 2-aminobenzoate, quinoline

Bak and Widdel (1986a)

Desulfobacterium phenolicumd

Phenol, p-cresol, benzoate, phenylacetate, indole, 4-hydroxyphenylacetate, 2-hydroxybenzoate, 4-hydroxybenzoate, phenylalanine, 2-aminobenzoate

Bak and Widdel (1986b)

Desulfobacterium catecholicume

Catechol, resorcinol, 4-hydroxybenzoate, hydroquinone, benzoate 2-aminobenzoate, protocatechuate, phloroglucinol, pyrogallol

Szewzyk and Pfennig (1987)

Desulfobacterium anilinic

Aniline, 2-aminobenzoate, 4-aminobenzoate, indolylacetate, quinoline

Schnell et al. (1989)

strain Cat2c

Phenol, catechol, m-cresol, p-cresol, benzoate, phenylacetate, phenylpropionate 4-hydroxybenzoate, 3,4-dihydroxybenzoate, phenylalanine

Schnell et al. (1989)

strain SAXc

Benzoate, phydroxybenzoate, phenol, phenylacetate, phenylalanine

Drzyzga et al. (1993)

Desulfotomaculum Catechol, phenol, m-cresol,, Kuever et al. strain Grollf p-cresol, benzoate, (1993) 3-hydroxybenzoate, benzaldehyde benzyl alcohol, phenylacetate, phenylpropionate Desulfovibrio inopinatus

Hydroxyhydroquinone (1,2,4-trihydroxybenzene)

Reichenbecher and Schink (1997)

Aromatic hydrocarbons ‘‘Desulfobacula toluolica’’

Toluene, p-cresol, benzaldehyde, benzoate, phenylacetate, p-hydroxybenzoate p-hydroxybenzaldehyde

Rabus et al. (1993)

355

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

. Table 9.5 (continued) Aromatic substratea

Reference

Toluene, p-cresol, benzaldehyde, benzoate, phenylacetate, phenylpropionate p-hydroxybenzoate

Beller et al. (1996)

strain mXyS1c

Toluene, m-xylene, m-ethyltoluene, m-isopropyltoluene, benzoate, m-methylbenzoate

Harms et al. (1999)

strain oXyS1h

Toluene, o-xylene, o-ethyltoluene, o-methylbenzyl alcohol benzoate, o-methylbenzoate, benzylsuccinate

Harms et al. (1999)

strain NaphS2h

Naphthalene, 2-naphthoate, benzoate

Galushko et al. (1999)

Organism strain PRTOL1

g

a Information about additional aromatic substrates is provided in the respective references b Has to be reclassified as Desulfobacterium niacini (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 classified 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)

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 1989; Heider and Fuchs 1997a, b; Harwood et al. 1999). Important principles of aromatic compound degradation recognized in the nonsulfate-reducing 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 1989; Lack and Fuchs 1992, 1994), the involvement of an a-oxidation reaction in the

conversion of phenylacetate to benzoyl-CoA (Mohamed et al. 1993), and the reductive removal of hydroxyl groups (Tschech and Schink 1986; Gibson et al. 1997). Free benzoate is simply activated to benzoyl-CoA in an ATP-consuming reaction (Geissler et al. 1988; Altenschmidt et al. 1991). Some 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 4-aminobenzoate, which via ligation with acetyl-CoA 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 p-cresol 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 methyl-group oxidation to 3-hydroxybenzoate because the latter compound was detected in m-cresol-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 section > ‘‘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 (Mu¨ller 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-1-carboxyl-CoA (Boll and Fuchs 1995, 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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 sulfatereducing bacteria are nicotinate (Imhoff-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 6-hydroxynicotinate 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 (Gribc-Galic and Vogel 1987) and 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 > Eq. 9.18, as demonstrated by measurement of the degradation balance. Another toluenedegrading sulfate reducer, strain PRTOL1, was isolated from fuel-contaminated subsurface soil (Beller et al. 1996). C6 H5 CH3 þ 4:5SO4 2 þ 2Hþ þ 3H2 O ! 7HCO3 þ4:5H2 S DG 0 ¼
205 kJ=mol toluene ð9:18Þ 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-xylene-degrading 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 utilize

9

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 1995; 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 denitrifier (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. Fumarate-dependent 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. 9.16). Further degradation of benzylsuccinate is proposed to proceed via reactions analogous to b-oxidation of a-methyl-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 sulfate-reducing 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, naphthalene-degrading strain NaphS2 is able to grow on 2-naphthoate, but not on 1-naphthoate (Galushko et al. 1999). A different initial mechanism of anaerobic naphthalene 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

357

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

COO− COO− COO− CH3

COSCoA

Fumarate COO−

Toluene

Benzylsuccinate

Benzoyl-CoA

. Fig. 9.16 Anaerobic, fumarate-dependent activation of toluene to benzylsuccinate in Desulfobacula toluolica and strain PRTOL1. Further degradation of benzylsuccinate to the central intermediate benzoyl-CoA is not completely understood

and the fact that aerobic bacteria initiate alkane activation always with O2 as cosubstrate (monooxygenase 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 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 (airexcluded) 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 9.6. Anaerobic utilization of various long-chain 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. 9.17) 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 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. 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 alkanedegrading 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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

9

. Table 9.6 Sulfate-reducing bacteria with the capacity to use aliphatic hydrocarbons as growth substrates n-Alkanes

1-Alkenes

Reference

b

28–30

C12–C20

C14, C16, C18

Aeckersberg et al. (1991)

c

C14, C16, C18

Organism Hxd3 Pnd3

Aliphatic hydrocarbon utilizeda

Optimum temperature ( C) 30

C14–C17

TD3cd

55–65

C6–C16, 3-methyloctane

AK01e

26–28

C13–C18

Aeckersberg et al. (1994, 1998) Rueter et al. (1994), Ehrenreich (1996)

C15, C16

So and Young (1999a)

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

∆G (kJ/mol SO42–)

–30

gaseous

liquid

–40

–50

–60

–70 2

4

6

8

10

12

14

16

C-Atoms per chain (n)

. Fig. 9.17 Free energy change of sulfate reduction with n-alkanes 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)/4SO42
! nHCO3
+ (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)

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, 13 12 C/ C analyses are in favor of an anaerobic methane oxidation. Residual methane in the zone of its anaerobic depletion is 13 C-enriched (and 2H-enriched), 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 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, 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 · L 1 · day 1 or were even lower. However, at a gas seep, volumetric sulfate-reduction rates as high as 2.5 mmol · L 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 14CO2 during methanogenesis but no net oxidation of methane has been measured in cultures of methanogenic archaea (Zehnder and Brock 1979, 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, 14C-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.

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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 13 12 C/ C-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 sulfate-reducing bacteria, but rather by a group of archaea (eventually identified as methanogens) 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. 9.11). 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; E0 = 0.269 V at pH 7.5) to render methane oxidation thermodynamically feasible. þ

CH4 þ 3H2 O ! HCO3 þ 4H2 þ H DG ¼ 15:7 kJ=mol SO4 2 þ 4H2 þ Hþ ! HS þ 4H2 O DG ¼ 15:7 kJ=mol Sum :

CH4 þ SO4 2 ! HCO3 þ HS þ H2 O DG ¼

31:4 kJ=mol

ð9:19Þ ð9:20Þ ð9:21Þ

(calculated for 25  C; pH = 7.5; CH4 partial pressure = 105 Pa; H2 partial pressure = 0.12 Pa; SO42
concentration = 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. 1989) 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, namely, a maximum rate (Vmax) of 90 mol H2 g
1 · h
1 (see sections > ‘‘Overview of Principal Properties,’’ > ‘‘Sulfate-Reducing Bacteria and Archaea’’ in this chapter) and a half-saturation 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 1984b; 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, oxygendepleted 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, E00 =
0.41 V) in aqueous surroundings (according to 2Fe + 2H+ ! 2Fe2+ + 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 ! Fe2+ + 2e
) 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 (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

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 (E00 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 sulfate-reducing 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 allows 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 propionyl-CoA. 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 section > ‘‘Propionate’’ in this chapter Kremer and Hansen 1988; > Fig. 9.15). 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 (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 3,4,5trimethoxybenzoate 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 > Eq. 9.22: 2þ H3 N CH2 CH2 CH2 SO3 ! CH3 COO þ 2CO2 þ 2NH4 þ þ 2HS þ Hþ ð9:22Þ 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 > Eq. 9.23: 2 O3 S

CH2 CHðNH3 þ ÞCOO þ 2H2 O ! 2CH3 COO

þ 2CO2 þ SO4 2 þ HS þ 2NH4 þ þ Hþ ð9:23Þ Desulfovibrio species may grow with ethanol or lactate in the absence of sulfate if cocultured 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 H2-scavenging partner in the absence of sulfate, 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, E00 = 0.190 V; 2H+/H2, E00 = 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, E00 = 0.5 V) and acetyl phosphate to free acetate; the latter step allows substrate-level phosphorylation and generation of a proton gradient through ATP hydrolysis at the ATPase.

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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 propionateutilizing, syntrophic partners that were named as Syntrophobacter species turned out to be sulfate-reducing bacteria and members of the d-subclass (Do¨rner 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 propionate-oxidizing strains grew syntrophically by interspecies H2 transfer rather than by utilizing sulfate themselves. Apparently, the pathway of sulfate reduction is poorly developed in the propionate oxidizers.

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 acetylCoA 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 ! acetyl-CoA + PPi; in addition to succinyl-CoA: acetate transferase), phosphoenolpyruvate (PEP) synthetase (pyruvate + ATP ! PEP + AMP + Pi), and PEP carboxylase (PEP + HCO3 ! 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

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, b, c; 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, b, c) is explained by the pyruvate synthase reaction. Pyruvate synthase or pyruvate: ferredoxin oxidoreductase (PFOR) that carboxylates acetyl-CoA reductively (acetyl-CoA + CO2 + 2e + 2H+ ! 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

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 Stickland 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, b, c; Badziong et al. 1979; Brandis and Thauer 1981; Brysch et al. 1987). However, several newly isolated, completely oxidizing sulfatereducing bacteria grew with H2 plus CO2 (or formate) in the absence of other carbon compounds (Widdel 1980; Widdel and Pfennig 1981; Pfennig et al. 1981). Labeling studies with formate-utilizing Desulfoarculus (formerly Desulfovibrio) baarsii (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

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

than truly autotrophic sulfate-reducing bacteria; hence, direct enrichment of the latter (autotrophic bacteria) from natural samples under autotrophic conditions is unlikely. 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), namely, 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 COdehydrogenase 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) suggest 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 acetyl-CoA oxidation. Another incomplete oxidizer, Desulfomicrobium apsheronum, also has been reported to grow autotrophically (Rozanova et al. 1988).

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 (Riederer-Henderson 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 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).

9

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/L for growth (Stetter 1992). Phylogenetic analyses revealed that the genus Archaeoglobus represents a lineage within the Euryarchaeota (Woese et al. 1991) with particular relationships to methanogenic archaea; Archaeoglobus is unrelated to the sulfur-metabolizing and fermentative extreme thermophiles of the Crenarchaeota. An overview of physiological properties of sulfate-reducing Archaeoglobus species is given in > Table 9.7. 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).

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 sulfatereducing bacteria (Dahl and Tru¨per 1999b). The presence of the enzymatic activities essential for dissimilatory reduction of sulfate (ATP sulfurylase, APS reductase, and sulfite reductase) was demonstrated in A. fulgidus (Speich and Tru¨per 1988; Dahl et al. 1994). In > Table 9.3, the sulfite reductase from A. fulgidus is compared to 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, 1990). The coding gene for sulfate adenylyltransferase (sat) was cloned and found to exhibit homology with the coding genes of homo-oligomeric 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

363

364

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

. Table 9.7 Physiological properties of sulfate-reducing Archaeoglobus speciesa Organic substrates utilized with SO42
and/or S2O32


Species

Temp. opt. [ C] Formate Acetate Lactate Pyruvate Others

H2 utilization Lithoautotrophic Lithoheterotrophic With With (+acetate) with SO42
S2O32
SO42


A fulgidus strain VC-16b 83

+

nr

+

nr

Formamide, glucose, starch, peptone, methanol, ethanol




+

nr

A fulgidus strain Zb

75–80

+




+

+

2,3-Butandiol, fumarate




+

nr

A fulgidus strain 7342b

76







+

+

Valerate

c

A profundus strain AV18b

82

nr

+

A lithotrophicus

80

nr

nr







+

+

+

c







+ (obligate lithoheterotrophic)

nr

nr

+

nr

nr

c

Symbols: +, utilized;
, not utilized; nr, not reduced Species of the genus Archaeoglobus 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), and 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 a

160 kDa was determined, and the protein was found to contain one FAD and [FeS] clusters (Speich and Tru¨per 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 additional 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-[4Fe-4S]-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).

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 (CO-dehydrogenase-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 heptaprenyl 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 9.7. 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 (Mo¨ller-Zinkhan et al. 1989; Zellner et al. 1989b). Lactate is oxidized to acetylCoA via lactate dehydrogenase and pyruvate:ferredoxin oxidoreductase (PFOR; Mo¨ller-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 2+ D-lactate dehydrogenase activity and contained Zn 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 and a heterotetrameric (abgd)

9

structure and to contain thiamine pyrophosphate and ironsulfur clusters (Kunow et al. 1995). Further oxidation of acetylCoA 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. 9.11b). 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; Mo¨ller-Zinkhan et al. 1989; Mo¨ller-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 acetylCoA into a bound methyl group and bound CO, or the reverse reaction. The methyl carrier is tetrahydromethanopterin (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 gsubunit and parts of the d-subunit; 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-methyleneH4MPT reductase; N5,N10-methylene-H4MPT dehydrogenase; N5,N10-methenyl-H4MPT cyclohydrolase; formylmethanofuran: H4MPT formyltransferase; and formylmethanofuran dehydrogenase (Mo¨ller-Zinkhan 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 acetatedegrading methanogens (Schmitz et al. 1991; Klein et al. 1993; Schwo¨rer et al. 1993). Factor F420 serves as H2 acceptor for N5,N10-methylene-H4MPT reductase and N5,N10methylene-H4MPT dehydrogenase. The natural electron acceptor for formylmethanofuran dehydrogenase is unknown. oxidoreductase A membrane-bound F420H2:quinone complex was purified from A. fulgidus. This enzyme complex

365

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9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 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. Even though Archaeoglobus 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 (Mo¨llerZinkhan 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. Malate dehydrogenase 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 (Aale´n et al. 1997). Malate, Isocitrate, and Glutamate

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 ( Eq. 9.25. n S8 þ HS
8

! Snþ1 2
þ Hþ

ð9:25Þ

In media around a pH of 7, the predominant species of polysulfide are tetrasulfide (S42
) and pentasulfide (S52
). These two polysulfides interconvert rapidly, as shown in > Eq. 9.25, and are also in equilibrium with lower concentrations of other polysulfides. Therefore, it is unknown which of the two polysulfides is the preferred substrate for polysulfide reductase. PPi formed from sulfide and tetrathionate was shown to be reduced by formate-utilizing cells of W. succinogenes (Klimmek et al. 1991). The concentration of polysulfide in medium containing 1 mM sulfide (HS
+ H2S) at pH > 6 was shown to exceed 10 mM (Schauder and Mu¨ller 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.

367

368

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 sulfidescavenging 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 1995) 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 (Bu¨hler 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 deleyianum, 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 sulfurcontaining 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 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)Sud-His6 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. 9.18; Hedderich et al. 1999).

Polysulfide Reductase The enzyme that catalyzes the reduction of polysulfide sulfur to sulfide is termed ‘‘polysulfide reductase’’ (Psr). This 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 molybdenum-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.

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

OM

+ + +

Periplasm

H2

HydB Ni

HydA 2 e– Fe/S

2 H+

CM

– – –

9

Cytoplasm

HydC Cytb 2 e–

H+ + [S] Sud HS–

H+ Mo PsrA

2 e– Fe/S

MKH– MK

PsrB

PsrC

H+

1 / ATP 3

+ + +

– – –

1 / ADP 3 + Pi

. Fig. 9.18 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 are assumed to be required for electron transfer. The mechanism for generation of a proton gradient is not known. Possibly, protons are translocated via protein-bound 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))

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 is used as electron acceptors (Schro¨der 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, b). 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 (Kro¨ger 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 diffusion and collision was suggested. Collision of hydrogenase or formate dehydrogenase, respectively, with the polysulfide reductase is regarded as

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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 transmembranespanning 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 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. 9.18). 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 (Kro¨ger et al. 1980). The substrate-binding sites of hydrogenase and of formate dehydrogenase both face the periplasm (Kro¨ger 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).

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 9.2).

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 Kro¨ger et al. 1988; Thauer 1988, 1989; Thauer et al. 1989). [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. 9.12). 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. 9.12c). 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)+ (E00 = 0.32 V) with malate (E00 = 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 (Kro¨ger et al. 1988). A comparison of the modifications of the citricacid cycle found in sulfate- and sulfur-reducing bacteria is presented in > Fig. 9.12. Inasmuch as neither Desulfuromonas nor Desulfurella gains 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 isocitrate, to the postulated electron donor for the sulfur reductase (DE0 around 0.2 V) could pump 2H+/2e or 4H+/acetate. As in Desulfobacter, succinate oxidation in Desulfuromonas acetoxidans to fumarate (E00 = +0.033 V) with menaquinone (E00 = 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, E0 = 0.19 V for 10 mM H2S; > Fig. 9.14b). It is more likely that electron transport from succinate (oxidation) to sulfur (reduction) with a redox span of DE0 = 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 h (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

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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 (Kro¨ger et al. 1988). Indeed, the cytochrome c551∧.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 c551∧.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 c551∧.5 was cloned and heterologously overexpressed in Desulfovibrio desulfuricans. The purified recombinant cytochrome c551∧.5 had the same biochemical and metal-reducing properties as the protein from Desulfuromonas acetoxidans (Aubert et al. 1998b). Structural analysis revealed strong analogies between the triheme cytochrome c551∧.5 and the tetraheme cytochrome c3. The region that harbors the heme II group in c3 is not present in c551∧.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 substratelevel 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’’ sulfur-reducing archaea, as far as their biochemistry has been investigated. In addition, results from investigations on Pyrococcus furiosus are included;

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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 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 ‘‘H2-evolving’’ 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 enzyme 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 ad-subunits 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 a-subunit (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 (Scha¨fer and Scho¨nheit 1992; 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 1983). However, growth due to this reaction has not been demonstrated.

Metabolism of Organic Electron Donors Carbohydrates

Most of the archaeal sulfur reducers 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; Scho¨nheit and Scha¨fer 1995; Kengen et al. 1996; Hedderich et al. 1999). The thermoacidophilic, sulfur-reducing archaeon Thermoproteus tenax utilizes, besides other substrates, glucose for growth by sulfur respiration (Zillig et al. 1981; Fischer et al. 1983). Part of the glucose can be transiently stored as glycogen (Ko¨nig et al. 1982). Glucose in the energy metabolism is completely oxidized (Selig and Scho¨nheit 1994). Labeling experiments with [13C]- and [14C]-glucose and enzymatic studies demonstrated that T. tenax employs in parallel a modified Embden-Meyerhof-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 (Scho¨nheit and Scha¨fer 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-2,6-bisphosphate, the classical effectors of ATP-PFK (Siebers et al. 1998). Phylogenetic analysis of the PPi-PFK encoding

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

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 glucose oxidation (Siebers et al. 1997). Decarboxylation of pyruvate to acetyl-CoA is catalyzed by pyruvate:ferredoxin oxidoreductase (Selig and Scho¨nheit 1994; Scho¨nheit and Scha¨fer 1995). This enzyme is also operative in Pyrobaculum islandicum and Pyrococcus furiosus (Scha¨fer and Scho¨nheit 1991). Further oxidation of acetate to CO2 in T. tenax and P. islandicum involves the citric-acid cycle (Selig and Scho¨nheit 1994). The sugar metabolism in Pyrococcus furiosus, which has been intensively investigated, has many parallels to that in Thermoproteus tenax (Mukund and Adams 1991; Scha¨fer and Scho¨nheit 1992; Kengen et al. 1994, 1996; Scha¨fer 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 (Scho¨nheit and Scha¨fer 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 Scho¨nheit 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, ribulose-1,5-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 (Scha¨fer 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 acetyl-CoA carboxylase and propionyl-CoA carboxylase function as the main CO2-fixation enzymes. A 3-hydroxypropionate cycle is proposed for these organisms as route of CO2 fixation (Menendez et al. 1999), where two moieties of CO2 are fixed by the aforementioned enzymes, and glyoxylate is formed for further synthesis of organic compounds from malyl-CoA, while acetyl-CoA is concomitantly

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regenerated. This pathway has originally been detected in the phototrophic bacterium Chloroflexus aurantiacus (Holo 1989; Strauss et 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 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

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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 endogenous hydrogen donors in the dark (Tru¨per 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 Desulfurospirillum deleyianum (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 ( Table 9.8.

Physiological and Practical Prerequisites for Genetic Studies in Sulfate-Reducing Bacteria Studies aiming at genetic manipulation of sulfate-reducing bacteria have mostly been carried out thus far with Desulfovibrio desulfuricans and D. vulgaris strains because various proteins in these species have been studied in great detail and because these species can be cultivated relatively easily. Other practical reasons for choosing these species were high plating efficiencies and antibiotic sensitivities for potential selection of mutants (Voordouw and Wall 1993; van Dongen et al. 1994). In the application of plating techniques for sulfate-reducing bacteria, anoxic growth conditions have to be guaranteed. van den Berg et al. (1989) developed a plating technique with a recovery of 50–100 %. Aerobically prepared agar plates were stored under an anoxic atmosphere until use. Desulfovibrio vulgaris was then plated under oxic conditions in an agar overlay of 3 mL. Immediately after solidification of the agar, the plates were incubated under argon (Ar)/CO2 in the presence of Na2S2O4 as O2 scavenger in the incubation container. Hence, anoxic conditions were necessary during growth, but not during plating. The presence of alternative modes of energy conservation is important for genetic studies of catabolic capacities. Desulfovibrio species can gain energy either from a respiratory metabolism with H2 or organic electron donors and with sulfate or sometimes nitrate as electron acceptors (> Table 9.1), or from fermentative degradation of pyruvate. This allows the genetic inactivation of genes involved in one process, while other processes are still operative to sustain viability. A mutant of Desulfovibrio desulfuricans was generated by chemical mutagenesis that could no longer use H2 for the reduction of sulfate, but could still grow with lactate and sulfate (Odom and Wall 1987). The use of DNA inserts with resistance markers and application of antibiotics is a common method of generating and selecting mutants. Even though sulfate-reducing bacteria are generally resistant to many antibiotics (Saleh et al. 1964), some antibiotics proved to be useful. Plasmid-borne resistance to streptomycin and sulfonamides was expressed in Desulfovibrio desulfuricans (Powell et al. 1989) and resistance to chloramphenicol in D. vulgaris (van den Berg et al. 1989).

9

Delivery Systems for DNA A prerequisite for defined genetic manipulation is the availability of tools to deliver genetic information into cells. All tools commonly used in genetic studies of other microorganisms have also been applied to sulfate-reducing bacteria.

Transduction Knowledge about the occurrence of bacteriophages in sulfatereducing bacteria and their applicability for transduction is limited. Handley et al. (1973) presented first evidence for bacteriophages in sulfate-reducing bacteria. They observed bacteriophage-like particles in cultures of Desulfovibrio vulgaris that had been treated with mitomycin C. A defective bacteriophage, termed Dd1, was demonstrated to mediate transduction in Desulfovibrio desulfuricans strain ATCC (Rapp and Wall 1987). Mixing and incubating D. desulfuricans strains resistant to either rifampicin or nalidixic acids resulted in the development of colonies simultaneously resistant to both antibiotics at higher rates than expected for spontaneous mutations. The phage Dd1 was identified as the transducing agent of the resistance markers and found to resemble morphologically the coliphages T7 and T3. A practical limitation to the use of bacteriophage Dd1 is its inability to transfer resistance to other strains of Desulfovibrio desulfuricans or to other Desulfovibrio species (Voordouw and Wall 1993). A different bacteriophage was isolated from marine sediment that could lyse cells of Desulfovibrio salexigens, as demonstrated by plaque formation. Morphological and molecular characteristics suggested a relation of the isolate to the lysogenic bacteriophage l, but the potential of this phage for genetic transfer was not investigated (Kamimura and Araki 1989). Treatment with UV light allowed the induction of lytic bacteriophages in Desulfovibrio vulgaris (Hildenborough). Subsequent mapping studies indicated the presence of two prophages in D. vulgaris (Seyedirashti et al. 1991, 1992).

Conjugation Conjugation has been used several times to transfer broad host-range plasmids belonging to the incompatibility group Q (IncQ) from Escherichia coli to Desulfovibrio species. Derivatives of plasmid pSUP104 were transferred with a frequency of about 10 2 from Escherichia coli to Desulfovibrio vulgaris. Stable maintenance of the plasmids in D. vulgaris could be demonstrated (van den Berg et al. 1989). At the same time, Powell et al. (1989) reported on a retrotransfer of plasmid R300B from Desulfovibrio back to E. coli. The cytochrome c3-encoding cyc gene from Desulfovibrio vulgaris (Hildenborough) was ligated into plasmid pJRDC800-1, transferred by conjugation from E. coli to Desulfovibrio desulfuricans G200, and then functionally expressed in the new host

375

376

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

. Table 9.8 Genes cloned and characterized from sulfate-reducing bacteriaa Gene(s) product

Gene(s) designation Organismb

References

[Fe]hydrogenase

hydA, hydB

D. vulgaris (Hildenborough)

Voordouw and Brenner (1985)

D. vulgaris subsp. oxamicus (Monticello)

Voordouw et al. (1989a)

D. desulfuricans

Hatchikian et al. (1999)

D. fructosovorans

Casalot et al. (1998) Stokkermans et al. (1989)

Voordouw et al. (1985)

Protein, unknown function

hydC

D. vulgaris (Hildenborough)

[NiFe]hydrogenase

hynA, hynB

D. gigas

Voordouw et al. (1989a) Li et al. (1987) Voordouw et al. (1989b)

[NiFeSe]hydrogenase

D. vulgaris (Miyazaki F)

Deckers et al. (1990)

D. fructosovorans

Rousset et al. (1990)

hynC

D. fructosovorans

Rousset et al. (1993)

hysA, hysB

Desulfomicrobium baculatum

Menon et al. (1987), (1988) Voordouw et al. (1989b)

NADP-reducing hydrogenase

hndA, B, C, D

D. fructosovorans

Malki et al. (1995)

Cytochrome c3

cyc

D. vulgaris (Hildenborough)

Voordouw and Brenner (1986)

CycD

D. desulfuricans (Norway)

Aubert et al. (1997)

Acidic cytochrome c3

D. africanus

Magro et al. (1997)

Basic cytochrome c3

D. africanus

Magro et al. (1997)

Cytochrome c

Desulfomonile tiedjei (DCB-1) Louie et al. (1997) D. vulgaris (Hildenborough)

Cytochrome c553

van Rooijen et al. (1989)

High-molecular-mass cytochrome c (Hmc)

hmc

D. vulgaris (Hildenborough)

Pollock et al. (1991)

Nonaheme cytochrome c

ddE

D. desulfuricans (Essex 6)

Fritz (1999)

The hmc operon, potential transmembrane redox hmc, Orf2–6, Rrf1–2 protein complex

D. vulgaris (Hildenborough)

Rossi et al. (1993)

Flavodoxin

D. vulgaris (Hildenborough)

Voordouw (1988a)

fla

Curley and Voordouw (1988) Krey et al. (1988) Carr et al. (1990) D. vulgaris (Miyazaki F)

Ferredoxin Rubredoxin

rub

Kitamura et al. (1998)

D. salexigens

Helms et al. (1990)

D. desulfuricans

Helms and Swenson (1991)

D. gigas

Chen et al. (1994b)

D. vulgaris (Hildenborough)

Voordouw (1988a) Brumlik and Voordouw (1989)

D. vulgaris (Miyazaki F)

Kitamura et al. (1997)

Desulfoarculus baarsii

Pianzzola et al. (1996)

D. vulgaris (Hildenborough)

Brumlik et al. (1989)

Desulfoarculus baarsii

Pianzzola et al. (1996)

Rubredoxin oxidoreductase

rbo

Desulforedoxin

dsr

D. gigas

Brumlik and Voordouw (1990)

OMPase (orotidine-50 -phosphate decarboxylase)

pyrF

D. vulgaris (Hildenborough)

Li et al. (1986)

Nitrogenase, Fe protein

nifH

D. gigas

Postgate et al. (1988)

DdeI, restriction endonuclease, methylase

hsdM, hsdR

D. desulfuricans

Howard et al. (1986)

Kent et al. (1989) Sznyter et al. (1987)

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

9

. Table 9.8 (continued) Gene(s) product

Gene(s) designation Organismb

References

Rubrerythrin

rbr

D. vulgaris (Hildenborough)

Prickril et al. (1991)

Methyl-accepting chemotaxis proteins, DcrA-L

dcrA-L

D. vulgaris (Hildenborough)

Dolla et al. (1992) Deckers and Voordouw (1994a) Deckers and Voordouw (1996)

APS reductase

aprBA

D. desulfuricans (Essex)

Fritz (1999)

Assimilatory sulfite reductase

asr

D. vulgaris (Hildenborough)

Tan et al. (1991)

D. vulgaris (Miyazaki F)

Kitamura et al. (1995)

selC

Desulfomicrobium baculatum

Tormay et al. (1994)

MOP, molybdenum-containing aldehyde oxidoreductase,

MOP gene

D. gigas

Thoenes et al. (1994)

FOR, pyruvate-ferredoxin oxidoreductase

por

D. africanus

Pieulle et al. (1997)

D. desulfuricans (Norway)

Fons et al. (1987)

Cytochrome c oxidase-like protein Selenocysteine-inserting tRNA (tRNA

Sec

)

Proline and leucine biosynthesis a

This table has been modified from Voordouw (1993) The genus Desulfovibrio is abbreviated with D.

b

(Voordouw et al. 1990). Interestingly, broad host-range plasmids from the incompatibility group P and W (IncP and IncW) could not be transferred to D. desulfuricans G100A (Argyle et al. 1992). This finding pointed at the specificities in the ability of Desulfovibrio to receive or maintain broad host-range plasmids.

Endogenous Plasmids The presence of plasmids in several Desulfovibrio species was previously reported (Postgate et al. 1984, 1986). Desulfovibrio gigas (NCIMB 9332) carries two plasmids of the sizes 105 and 60 kb, and D. vulgaris (Hildenborough; NCIMB 8303) a single plasmid of 195 kb. D. desulfuricans strain Berre sol and D. vulgaris strain Wandle both carry a single plasmid. No plasmids could be detected in 10 other Desulfovibrio species, including D. salexigens and D. africanus. A small 2.3 kb plasmid, designated pBG1, was isolated from Desulfovibrio desulfuricans strain G100A and sequenced (Wall et al. 1993). Plasmid pBG1 was present in about 20 copies per genome. This plasmid replicates in D. desulfuricans strain G100A and D. fructosovorans, but not in Escherichia coli. The analysis of the sequence of plasmid pBG1 allowed a replicon area to be assigned to a sequence. Integration of pBG1 fragments that included this small replicon into derivates of the IncQ plasmid RSF1010 generated composite plasmids that were stable and replicated in E. coli, D. desulfuricans strain G100A, and D. fructosovorans. Several recombinant plasmids that are most effective in Desulfovibrio were constructed that carried genes for resistance to antibiotics, e.g., chloramphenicol. These plasmids could be transferred by either electroporation or conjugation (Rousset et al. 1998a).

Transformation The first successful transformation of a sulfate-reducing bacterium was reported by Rousset et al. (1991). A recombinant plasmid belonging to the IncQ group was transformed into Desulfovibrio fructosovorans by means of electroporation (Dower et al. 1988). Transformation via electroporation has also been successfully applied to Desulfovibrio desulfuricans (Aubert et al. 1998b).

Creation of Mutants Stability and function of recombinant DNA introduced into Desulfovibrio species or other sulfate-reducing bacteria could be hampered in principle by an endogenous restriction/modification system. Only little is known about these systems in sulfate-reducing bacteria. Two restriction endonucleases, DdeI and DdeII, were discovered in D. desulfuricans (Norway). Genes coding for DdeI endonuclease and methylase were identified (Howard et al. 1986; Sznyter et al. 1987).

Chemical Mutagenesis Mutants of Desulfovibrio desulfuricans strain ATCC 27774 that were deficient in H2 utilization but could still grow on lactate were generated by exposure to UV light (Odom and Wall 1987).

Transposon Mutagenesis Transposons can insert principally near or in any gene on the chromosome and completely annihilate gene function as a result

377

378

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

of gene disruption (Maloy et al. 1996). Wall et al. (1996) developed a transposon-based method for random mutagenesis in Desulfovibrio desulfuricans strain G20. No evidence was found for transposition of wild-type transposon Tn5, Tn9, and Tn10 in this sulfate reducer. The transposon Tn7 was found to insert into the genome of D. desulfuricans (G20) with a frequency of 10 4 to 10 3. The transposon Tn7 is target specific and inserts specifically into the attTn7 site. This directed insertion is mediated by the Tn7-encoded TnsD protein (Craig 1991; Bainton et al. 1993). Southern blot analysis demonstrated that transposon Tn7 inserted also in D. desulfuricans strain G20, specifically into the attTn7 site. By inactivation of the tnsD gene, Wall et al. (1996) were able to create a mutated Tn7, designated Tn7-IN1, that transposed randomly in the chromosome at a frequency of 10 6. An improved version of the transposon, designated Tn7K-IN1, carried a kanamycin-resistance cassette and mediated a Knr-phenotype after transposition. This transposon as part of gene constructs allows the integration of single copies of recombinant genes at neutral spots in the chromosome. Plasmids used for the delivery of Tn7, its derivatives, and adjacent genes by conjugation are not maintained in D. desulfuricans. The insertion element ISD1, which was discovered in D. vulgaris strain Hildenborough, is the first described transposable element detected in sulfate-reducing bacteria (Fu and Voordouw 1998). This element is 1.2 kb in length, encodes a transposase, and can actively transpose into different sites on the genome of D. vulgaris strain Hildenborough. Sequence analysis revealed a relation of ISD1 to the IS3 family. Members of the IS3 family have been isolated from Gram-negative and Grampositive bacteria (Fayet et al. 1990). ISD1 may be used as a platform to generate artificial transposons for random mutagenesis in Desulfovibrio species.

Gene Deletion Deletion of the hydN gene (coding for the [NiFe]-hydrogenase) in Desulfovibrio fructosovorans by marker-exchange mutagenesis was the first report of a successful gene replacement in sulfate-reducing bacteria (Rousset et al. 1991). The cloned hydN gene was replaced by a kanamycinresistance cassette, while the conserved flanking regions were maintained to allow homologous recombination. In the applied procedure, an IncQ plasmid was used as shuttle vector and transferred to D. fructosovorans by electroporation. Replacement of the chromosomal hydN gene with the marker was verified by expression of kanamycin resistance and a 90 % decrease of hydrogenase activity. Furthermore, analysis by southern hybridization confirmed the gene replacement. In subsequent studies, single and double mutants of the NADP+-reducing hydrogenase were generated in D. fructosovorans by this method of marker-exchange mutagenesis (Malki et al. 1997).

Fu and Voordouw (1997) developed a method for gene replacement based on a suicide plasmid in Desulfovibrio vulgaris strain Hildenborough to study the oxygen sensor DcrA. A suicide-integration plasmid (pDDcrA2CTB) was constructed from an IncQ plasmid carrying the cloned dcrA gene. The vector carried a dcrA allele disrupted by the cat gene (conferring resistance to chloramphenicol) and contained the counterselectable marker sacB (coding for levansucrase; Gay et al. 1983) from Bacillus subtilis. Plasmid pDDcrA2CTB was transferred from E. coli strain S17-1 to D. vulgaris by conjugation. Integration of plasmid pDDcrA2CTB into the chromosomal dcrA gene by the first event of homologous recombination was selected for by the presence of chloramphenicol. Addition of sucrose to the growth medium then selected for the second homologous recombination which resulted in excision of the plasmid from the chromosome and the replacement of the dcrA gene. In the presence of sucrose, the gene product of sacB is toxic for E. coli and other Gram-negative bacteria (Gay et al. 1983) and has therefore widely been used as a counterselectable marker for the rare second recombination event which yields clones cured from plasmid (Ried and Collmer 1987; Blomfield et al. 1991). The genes of the hmc operon (coding for the high-molecularmass cytochrome redox protein complex, the Hmc complex) and the rbo gene (like dcrA related to oxygen sensitivity), both of D. vulgaris, were also deleted employing the aforementioned method (Keon et al. 1997; Voordouw and Voordouw 1998).

Site-Directed Mutagenesis Site-directed mutagenesis was applied to study the signalpeptide consensus box of [NiFe] hydrogenase of Desulfovibrio vulgaris (Hildenborough) in a fusion protein with b-lactamase from E. coli (Nivie`re et al. 1992). Exchange of an arginine residue in the consensus box for a glutamate prevented export of the fusion protein from the cytoplasm of E. coli.

Sulfur-Reducing Bacteria Genome Sizes, Genomic Libraries, and Cloning of Genes The genome sizes of Desulfurella acetivorans and D. multipotens were reported to be around 1.9 Mb (Pradella et al. 1998). Genomic libraries of W. succinogenes were constructed using the bacteriophage EMBL-3 (Frischauf et al. 1983; Lauterbach et al. 1987). From these, subcloning of several genes was possible. These were frd genes coding for fumarate reductase (Lauterbach et al. 1987), fdh genes coding for formate dehydrogenase (Bokranz et al. 1991), psr genes coding for polysulfide reductase (Krafft et al. 1992), and sud gene coding for the periplasmic sulfide dehydrogenase (Kreis-Kleinschmidt et al. 1995).

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Physiological and Practical Prerequisites for Genetic Studies in Sulfur-Reducing Bacteria The sulfur-reducing bacterium Wolinella succinogenes can grow on agar plates when anoxic conditions are maintained. In addition, antibiotics can be used as selection markers, because W. succinogenes is sensitive to, for instance, kanamycin. Transformation is accomplished by means of electroporation and has to be performed under anoxic conditions like plating. The broad host-range plasmid pBR322 (Bolivar et al. 1977), which is commonly used as shuttle vector in Gram-negative bacteria (Maloy et al. 1996), can be used to transfer recombinant DNA from E. coli back to W. succinogenes.

. Table 9.9 Present status (June 2012) of families embracing (though not exclusively) dissimilatory sulfate- and sulfur-reducing prokaryotes. Genera in bold were mentioned in > Table 9.1 (Morphological and physiological properties of the genera of sulfate-reducing bacteria and archaea) and > Table 9.2 (Morphological and physiological properties of Bacteria and Archaea capable of respiratory reduction of elemental sulfur) of the publication by Rabus et al. (2006). Source: http://www.bacterio.cict.fr/classifgenerafamilies.html Spalte1 Genus From

> Table

Reference 9.1

Bacteria Family

Gene Deletion

Desulfarculaceae Desulfarculus

Family

For gene deletion, plasmid-hosted genes from W. succinogenes are disrupted by antibiotic-resistance cassettes (e.g., kanamycin) leaving homologous regions to both sides of the marker. Deletion of the chromosomal target genes is then accomplished by homologous recombination. Several genes have been deleted in W. succinogenes by this procedure: psr genes coding for polysulfide reductase (Krafft et al. 1995), the sud gene coding for periplasmic sulfide dehydrogenase (Kotzian et al. 1996), fdh genes coding for formate dehydrogenase (Lenger et al. 1997), frd genes coding for fumarate reductase (Simon et al. 1998), and hyd genes coding for hydrogenases (Gross et al. 1998a, b).

Desulfobacteraceae Desulfatibacillum

Cravo-Laureau et al. (2004)

Desulfatiferula

Cravo-Laureau et al. (2007)

Desulfatirhabdium

Balk et al. (2008)

Desulfoluna

Suzuki et al. (2008)

Desulfomusa

Finster et al. (2001)

Desulforegula

Rees et al. (2001)

Desulfosalsimonas

Jkeldsene al. (2010)

Desulfotignum

Kuever et al. (2001)

Desulfobacter Desulfobacterium Desulfobacula Desulfobotulus Desulfocella

Sulfur-Reducing Archaea At present, only initial steps in the establishment of systems for genetic manipulation of sulfur-reducing archaea have been undertaken. In contrast, a variety of genetic tools have already been developed for halophilic archaea (e.g., Cline and Doolittle 1987). Nevertheless, various genetic elements have been discovered in archaeal sulfur reducers, and new composite shuttle vectors are being developed. Several types of viruses were discovered in Thermoproteus species. A plasmid, termed pDL10, was found to be present in Desulfurolobus species (Zillig et al. 1996). Plasmid pGT5 was isolated from Pyrococcus abyssi (Erauso et al. 1996). A mobile intron from Desulfurococcus mobilis could be transferred to and established in Sulfolobus acidocaldarius (Aagaard et al. 1995). A new hybrid shuttle vector, designated pAG1, was constructed by combining portions of the archaeal plasmid pGT5 with the bacterial plasmid pUC19. The plasmid pAG1 was stably maintained and propagated both in bacteria and archaea (Aravalli and Garrett 1997). A different strategy to create a new shuttle vector is to incorporate the mobile intron from Desulfurococcus mobile into the bacterial vector pUC18 (Aagaard et al. 1996, > Table 9.9).

9

Desulfococcus Desulfofaba Desulfofrigus DesulfonemaDesulfosarcina Desulfospira Family

Desulfobulbaceae Desulfopila

Suzuki et al. (2007)

Desulfurivibrio

Sorokin et al. (2008a)

Desulfobulbus Desulfocapsa Desulfofustis Desulforhopalus Desulfotalea Family

Desulfohalobiaceae Desulfonatronospira

Sorokin et al. (2008b)

Desulfonauticus

Audiffrin et al. (2003)

Desulfothermus

Kuever et al. (2005)

Desulfovermiculus

Belyakova et al. (2006)

Desulfohalobium Desulfonatronovibrio

379

380

9

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

. Table 9.9 (continued) Spalte1 Genus Family

. Table 9.9 (continued) Reference

Spalte1 Genus Family

Desulfomicrobiaceae Desulfomicrobium

Family

Desulfonatronaceae Desulfonatronum

Family

Thermodesulfatator

Moussard et al. (2004)

‘‘Nitrospiraceae’’

Desulfocurvus

Klouche et al. (2009)

Leptospirillum

Hippe (2000)

Desulfomonas

Moore et al. (1976) McOrist et al. (1995)

Nitrospira

Watson et al. (1986)

Thermodesulfovibrio Archaea Family

Desulfuromonadaceae

Archaeoglobaceae

Malonomonas

Dehning and Schink (1989)

Geoglobus

Kashefi et al. (2002)

Pelobacter

Schink and Pfennig (1982)

Ferroglobus

Hafenbradl et al. (1996)

Archaeoglobus From > Table 9.2 Bacteria

Desulfuromonas Family

Syntrophaceae

Desulfurobacteriaceae

Smithella

Liu et al. (1999)

Balnearium

Takai et al. (2003)

Syntrophus

Mountfort et al. (1984)

Thermovibrio

Huber et al. (2002)

Desulfurobacterium

Desulfobacca Family

Desulfomonile

Desulfurellaceae Desulfurella

Syntrophobacteraceae Desulfoglaeba

Davidova et al. (2006)

Desulfosoma

Baena et al. (2011)

Desulfovirga

Tanaka et al. (2000)

Syntrophobacter

Boone and Bryant (1980)

Desulfacinum

Hippea Family

Desulfuromonadaceae (see also > Table 9.1) Desulfuromonas

Family

Desulfurellaceae (see also 9.1)

> Table

Desulforhabdus

Desulfurella

Thermodesulforhabdus Family

Miroshnichenko etal. (2009)

Baton et al. (1989)

Desulfuromusa

Family

Caldimicrobium

Bilophila

Desulfovibrio

Family

Thermodesulfobacteriaceae

Thermodesulfobacterium

Desulfovibrionaceae

Lawsonia Family

Reference

Family

Peptococcaceae Cryptanaerobacter

Juteau et al. (2005)

Dehalobacter

Holliger et al. (1998)

Desulfitibacter

Nielsen et al. (2006)

Desulfitispora

Sorokin et al. (2010)

Desulfitobacterium

Desulfovibrionaceae (see also 9.1)

> Table

Desulfovibrio Family

Desulfomicrobiaceae (see also 9.1)

> Table

Desulfomicrobium Family

Peptococcaceae (see also 9.1)

Desulfonispora

Denger et al. (1999)

Desulfurispora

Kaksonen et al. (2007)

Pelotomaculum

Imachi et al. (2002)

Peptococcus

Kluyver and Van Niel (1936)

Aminiphilus

Diaz et al. (2007)

Sporotomaculum

Braumann et al. (1998)

Aminobacterium

Baena et al. (1998)

Syntrophobotulus

Friedrich et al. (1996)

Aminomonas

Baena et al. (1999a)

Thermincola

Sokolova et al. (2005)

Anaerobaculum

Rees et al. (1997)

Thermoterrabacterium

Slobodkin et al. (1997)

Cloacibacillus

Ganesan et al. (2008)

Jonquetella

Jumas-Bilak et al. (2007)

Pyramidobacter

Downe et al. (2009)

Synergistes

Allison et al. (1992)

Desulfitobacterium (> Table 9.2) Desulfosporosinus Desulfotomaculum

> Table

Desulfitobacterium Family

Utkin et al. (1994)

Synergistaceae

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Acknowledgements

. Table 9.9 (continued) Spalte1 Genus

Reference

Thermanaerovibrio

Baena et al. (1999b)

Thermovirga

Dahle and Birkeland (2006)

Dethiosulfovibrio Family

Peptococcaceae (see also > Table 9.1) Desulfitobacterium

Family

Campylobacteraceae Arcobacter

Vandamme et al. (1991)

Campylobacter

Seebald and Veron (1963)

Dehalospirillum

Scholz-Muramatsu et al. (1995)

Sulfurospirillum Family

Chrysiogenaceae Chrysiogenes

Macy et al. (1996)

Desulfurispira

Sorokin et al. (2010)

Desulfurispirillum

Sorokin et al. (2007)

Archaea Family

Desulfurococcaceae Aeropyrum

Sako et al. (1996)

Desulfurococcus

Zillig et al. (1982)

Ignicoccus

Huber et al. (2000)

Ignisphaera

Niederberger et al. (2006)

Staphylothermus

Fiala et al. (1986)

Sulfophobococcus

Hensel et al. (1997)

Thermosphaera

Huber et al. (1998)

Stetteria Thermodiscus Family

Pyrodictiaceae Hyperthermus

Zillig et al. (1991)

Pyrolobus

Blo¨chl et al. (1997)

Pyrodiction Family

Sulfolobaceae Desulfurolobus

Zillig et al. (1986)

Metallosphaera

Huber et al. (1989)

Sulfurisphaera

Kurosawa et al. (1998)

Sulfurococcus

Golovacheva et al. (1985)

Acidianus Stygiolobus Sulfolobus Family

Thermoprotaceae Caldivirga

Itoh et al. (1999)

Thermocladium

Itoh et al. (1998)

Vulcanisaeta

Itoh et al. (2002)

Pyrobaculum Thermoproteus

9

This work was supported by the Max-Planck-Gesellschaft and the Fonds der Chemischen Industrie. We thank Heribert Cypionka (Oldenburg), Christine Dahl (Bonn), Gu¨nter Fritz (Konstanz), Achim Kro¨ger (Frankfurt), Peter Kroneck (Konstanz), Bernhard Schink (Konstanz), and Rolf Thauer (Marburg) for providing informative material. We would very much appreciate any type of comments and further information that may help to improve future releases of this chapter. Comments and information may be sent via email to Dr. Rabus or to Dr. Widdel.

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Widdel F (1986) Growth of methanogenic bacteria in pure culture with 2-propanol and other alcohols as hydrogen donors. Appl Environ Microbiol 51:1056–1062 Widdel F (1987) New types of acetate-oxidizing, sulfate-reducing Desulfobacter species, D. hydrogenophilus sp. nov., D. latus sp. nov., and D. curvatus sp. nov. Arch Microbiol 148:286–291 Widdel F (1988) Microbiology and ecology of sulfate-and sulfur-reducing bacteria. In: Zehnder AJB (ed) Biology of anaerobic microorganisms. Wiley, New York, pp 469–585 Widdel F (1992) Microbial corrosion. In: Finn RK, Pra¨ve P, Schlingmann M, Crueger W, Esser K, Thauer R, Wagner F (eds) Biotechnology focus 3: fundamentals, applications, information. Hanser, Munich, pp 261–300 Widdel F, Bak F (1992) Gram-negative mesophilic sulfate-reducing bacteria. In: Balows A, Tru¨per HG, Dworkin M, Harder W, Schleifer K-H (eds) The prokaryotes, vol 3, 2nd edn. Springer, New York, pp 3352–3378 Widdel F, Pfennig N (1977) A new anaerobic, sporing, acetate-oxidizing, sulfatereducing bacterium, Desulfotomaculum (emend.) acetoxidans. Arch Microbiol 112:119–122 Widdel F, Pfennig N (1981a) Sporulation and further nutritional characteristics of Desulfotomaculum acetoxidans. Arch Microbiol 129:401–402 Widdel F, Pfennig N (1981b) Studies on dissimilatory sulfate-reducing bacteria that decompose fatty acids. I. solation of new sulfate-reducing bacteria enriched with acetate from saline environments. Description of Desulfobacter postgatei gen. nov., sp. nov. Arch Microbiol 129:395–400 Widdel F, Pfennig N (1982) Studies on dissimilatory sulfate-reducing bacteria that decompose fatty acids. II. Incomplete oxidation of propionate by Desulfobulbus propionicus gen. nov., sp. nov. Arch Microbiol 131:360–365 Widdel F, Pfennig N (1984) Dissimilatory sulfate-and sulfur-reducing bacteria. In: Krieg NR, Holt JG (eds) Bergey’s manual of systematic bacteriology, vol 1. Williams & Wilkins, Baltimore, pp 663–679 Widdel F, Kohring G-W, Mayer F (1983) Studies on dissimilatory sulfatereducing bacteria that decompose fatty acids. III. Characterization of the filamentous gliding Desulfonema limicola gen. nov., sp. nov., and Desulfonema magnum sp. nov. Arch Microbiol 134:286–294 Wight KM, Starkey RL (1945) Utilization of hydrogen by sulfate-reducing bacteria and its significance in anaerobic corrosion. J Bacteriol 50:238 Wilson LG, Bandurski RS (1958) Enzymatic reactions involving sulfate, sulfite, selenate, and molybdate. J Biol Chem 233:975–981 Woese CR (1987) Bacterial evolution. Microbiol Rev 51:221–271 Woese CR, Fox GE (1977) Phylogenetic structure of the prokaryotic domain: the primary kingdoms. Proc Natl Acad Sci USA 74:5088–5090 Woese CR, Magrum LJ, Fox GE (1978) Archaebacteria. J Mol Evol 11:245–252 Woese CR, Achenbach L, Rouviere P, Mandelco L (1991) Archaeal phylogeny: reexamination of the phylogenetic position of Archaeoglobus fulgidus in light of certain composition-induced artifacts. Syst Appl Microbiol 14:364–371 Wolfe RS, Pfennig N (1977) Reduction of sulfur by spirillum 5175 and syntrophism with Chlorobium. Appl Environ Microbiol 33:427–433 Wolfe BM, Lui SM, Cowan JA (1994) Desulfoviridin, a multimeric-dissimilatory sulfite reductase from Desulfovibrio vulgaris (Hildenborough). FEBS Lett 223:79–89 Wolin MJ, Wolin EA, Jacobs NJ (1961) Cytochrome-producing anaerobic vibrio, Vibrio succinogenes, sp. nov. J Bacteriol 81:911–917 Wood HG, Ragsdale SW, Pezacka E (1986) The acetyl-CoA pathway of autotrophic growth. FEMS Microbiol Rev 39:345–362 Woolfolk CA (1962) Reduction of inorganic compounds with molecular hydrogen by Micrococcus lactilyticus. J Bacteriol 84:659–668 Wu L-F, Mandrand MA (1993) Microbial hydrogenases: primary structure, classification, signatures and phylogeny. FEMS Microbiol Rev 104: 243–270 Yagi T (1969) Formate: cytochrome oxidoreductase of Desulfovibrio vulgaris. J Biochem 66:473–478 Yagi T (1979) Purification and properties of cytochrome c553, an electron acceptor for formate dehydrogenase of Desulfovibrio vulgaris, Miyazaki. Biochim Biophys Acta 548:96–105 Yamamoto I, Ishimoto M (1978) Hydrogen-dependent growth of Escherichia coli in anaerobic respiration and the presence of hydrogenases with different functions. J Biochem 84:673–679

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10 Denitrifying Prokaryotes James P. Shapleigh Department of Microbiology, Cornell University, Ithaca, NY, USA

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405 Defining the Denitrifiers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 406 Phylogenetic Distribution of Denitrification . . . . . . . . . . . . . . 407 Non-proteobacterial Denitrifiers . . . . . . . . . . . . . . . . . . . . . . . . 408 Horizontal Gene Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 414 Enzymology of Denitrification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 415 Regulation of Genes Required for Denitrification . . . . . . . . . 419 Environmental Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 421 Abstract Denitrification is the dissimilatory reduction of nitrate to nitrogen gas. This respiratory process requires four enzymes that produce three obligatory intermediates prior to production of the terminal product. Denitrification is found in diverse array of microbes including members of both bacteria and archaea. However, no bacterium has been described that solely depends on denitrification as a form of energy generation. All denitrifiers, with one exception, are aerobes. Genome sequencing has provided a better appreciation of the distribution of denitrification genes among microbes. Complete denitrification, the reduction of nitrate to N2, is less frequent than partial denitrification among sequenced bacteria. Partial denitrification chains of nearly all possible arrangments have been found. This includes chains with only a single enzyme or discontinuous chains of two or more enzymes. Nitrate reductase catalyzes the reduction of nitrate to nitrite and is used in a number of pathways other than denitrification; therefore, its distribution has not been a focus of this chapter. Nitrite reductase catalyzes the reduction of nitrite to nitric oxide and is the defining reaction of denitrification since it is the first step to produce a gaseous nitrogen oxide. There are two unrelated types of nitrite reductase, one of which has copper cofactors while the other contains heme-bound iron. The copper form has several different subtypes with N- and C-terminal extensions containing metal-binding sites. Some members of the Actinobacteria have a particularly large copper nitrite reductase with a membrane-bound domain of unknown function. Nitric oxide reductase catalyzes the reduction of nitric oxide to nitrous oxide. This enzyme is membrane bound and occurs in two subtypes referred to as cNor and qNor. The former receives electrons from cytochrome c while the latter carries an

N-terminal extension allowing it to oxidize quinol. Nitrous oxide reductase is a soluble copper-containing enzyme with one of the copper centers, designated the CuZ center, being unique to this enzyme. While most model denitrifiers use denitrification to support growth when oxygen is limiting, this may not be the case in all bacteria that contain genes encoding denitrification-associated nitrogen oxide reductases. Bacteria with partial chains consisting of a single enzyme may use that enzyme for alternative functions. For example, some Staphylococcus aureus subspecies aureus strains only contain nitric oxide reductase which is likely used for detoxification of nitric oxide. There are a number of bacteria which only contain nitrite reductase and the function of this enzyme is unclear in these organisms since its turnover will produce nitric oxide, which is toxic due to its reactivity with metal centers and other compounds. Environmental studies have found denitrification genes are nearly universal in environments that receive some exposure to oxygen. Quantitative studies have found that the genes for nitrous oxide reductase are frequently underrepresented compared to other denitrification genes. While common in soil and aquatic environments, denitrifiers are also found in association with humans. Sequencing of both skin and oral microbiomes has revealed a significant number of denitrifiers, consistent with the occurrence of both nitrate and nitrite in these areas.

Introduction One of the hallmarks of bacteria is the ability to use a wide variety of compounds as terminal oxidants for respiration. O2 (O2) respiration provides the most energy and so is the dominant form of respiration among living organisms. However, if O2 becomes limiting, many bacteria will continue respiration by switching the terminal electron acceptor to alternative compounds such as nitrate. Nitrate respiration occurs via two dissimilar pathways that utilize the same initial substrate but produce different end products (Canfield et al. 2010). One of these pathways, termed ammonification (or dissimilatory reduction of nitrate to ammonia [DNRA]), is carried out by bacteria such as Escherichia coli, and is marked by reduction of nitrate to nitrite and then to ammonia. The second pathway of nitrate respiration is denitrification, which is the reduction of nitrate to gaseous nitrogen oxides, and then nitrogen gas (> Fig. 10.1). The initial step in both denitrification and DNRA is the reduction of nitrate to nitrite (Richardson et al. 2001). The next step differentiates the two pathways. In DNRA, nitrite

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_71, # Springer-Verlag Berlin Heidelberg 2013

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Denitrifying Prokaryotes

NO3− (nitrate)

Nar Nap

NO2− (nitrite)

CuNir cd1-Nir

NO (nitric oxide)

cNor

Nos

N2O (nitrous oxide)

qNor

N2 (nitrogen)

. Fig. 10.1 The complete denitrification pathway. Chemical names for the nitrogen oxides are given in parentheses. Terms above and below arrows indicate the standard enzyme abbreviations used in the chapter

Denitrification

NO3−

N2

DN

fixa rog en

NO2−

RA DN

Nit

ion cat

tion

RA

undergoes a six electron reduction to ammonia (Simon 2002). In denitrification, nitrite is reduced to nitric oxide (NO), the defining step of this pathway (Zumft 1997). This conversion of a fixed, nongaseous, and biologically preferred form of nitrogen to a gaseous form is the reason this respiratory process is termed ‘‘denitrification.’’ The enzyme carrying out this reaction will be referred to as Nir. There are two types of Nir in denitrifiers. One contains copper and will be referred to as CuNir. The other contains heme and will be referred to as cd1-Nir. NO is not the terminal product of denitrification since it can be reduced to nitrous oxide (N2O) by a membrane-bound nitric oxide reductase, which will be referred to as Nor. As with Nir, there are two forms of Nor. One accepts electrons from quinol and will be referred to as qNor while the other accepts electrons from c-type cytochromes and will be referred to as cNor. The terminal step in denitrification is the reduction of N2O to nitrogen gas, N2, by nitrous oxide reductase, which will be referred to as Nos. The production of N2 connects denitrification to the nitrogen cycle via nitrogen fixation. While N2 is the terminal product of the pathway, it is important to note that it is not the case that a bacterium either has four enzymes of the denitrification pathway or none. Many bacteria have truncated or partial pathways that sometimes only include a single enzyme. The biologically mediated nitrogen cycle is shown in > Fig. 10.2. Nitrogen fixation produces ammonia. This ammonia, and the ammonia produced by DNRA, can be converted by nitrifying bacteria to nitrite and nitrate, substrates for denitrification. The anaerobic oxidation of ammonia, anammox, also results in the reduction of nitrogen oxides to N2 but is distinguished from denitrification by using electrons from ammonia (Martinez-Espinosa et al. 2011). Gayon and Dupetit carried out the first systematic study of nitrate conversion to gaseous forms of nitrogen in 1882 (Gayon and Dupetit 1882). Noting the loss of nitrate from decomposing sewage, they called it ‘‘denitrification’’ and were the first to isolate denitrifying bacteria (Gayon and Dupetit 1886), which they dubbed Bacterium denitrificans a and b. In the early stages of the study of denitrification, it was erroneously assumed that nitrate was releasing, and thus supplying, elemental O2 to organisms that subsequently carried out a reaction equivalent to O2 respiration. The observation of denitrification, although biologically significant, was unsettling to agronomists who soon realized that the addition of organic matter to soils could lead to the loss of fixed nitrogen. The agricultural importance of the process provided the impetus for much of the early work on denitrification, and by the end of the nineteenth century, denitrification had been reasonably well defined. Significant interest in the agricultural impact of denitrification has continued to be an important focus of research efforts.

ifi Nitr

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NH4

+

Assimilation

. Fig. 10.2 The biologically mediated nitrogen cycle. DNRA stands for dissimilatory reduction of nitrate to ammonia. Anammox stands for anaerobic ammonia oxidation

However, with the realization that NO and N2O play important roles in atmospheric and biological chemistry, research emphases have shifted to the environmental consequences of denitrification and the molecular mechanisms of enzymes and gene regulation (Brunekreef and Holgate 2002; Canfield et al. 2010; Ravishankara et al. 2009; Richardson et al. 2009).

Defining the Denitrifiers Denitrification is most frequently, but not always, used as a respiratory process. That is, reduction of nitrate and other intermediates proceeds through a series of protein complexes which results in the production of a proton motive force that can be used to generate ATP (van Spanning et al. 2007; Zumft 1997). Prokaryotes, mostly Bacteria, and a few Archaea, constitute the vast majority of organisms capable of denitrification. A number of fungal isolates carry out reduction of nitrate to N2O, but the contribution of this reduction to cell growth is variable (Kim et al. 2009; Nakanishi et al. 2010). More recent work has also found denitrification can occur in some multicellular eukaryotes (Risgaard-Petersen et al. 2006). With one or two exceptions, denitrifiers also respire O2 and, because it is usually available at higher concentrations and its reduction provides more energy to

Denitrifying Prokaryotes

the cell, O2 is the preferred electron acceptor. There are no bacteria known in which denitrification is the only means of producing ATP, meaning denitrification is never an essential physiological trait. The four separate enzymes catalyzing nitrate reduction to N2 produce freely diffusible products and catalyze reactions with a negative DG, making each step energetically favorable (Zumft 1997). This means each enzyme can potentially be an autonomous electron-accepting unit and, by extension, no reason a denitrifying bacterium must catalyze the complete reduction of nitrate to N2. While a few partial denitrifiers were found in the pre-genomic era (Denariaz et al. 1989; Greenberg and Becker 1977; Toffanin et al. 1996), genome sequencing has revealed partial denitrification is widespread. The occurrence of partial denitrification pathways makes defining a bacterium as a ‘‘denitrifier’’ somewhat tricky. Are denitrifiers only those bacteria that reduce nitrate to N2 or should the definition be more inclusive? For this chapter, a denitrifier is taken as any bacterium that has Nir, Nor, or Nos. Nitrate reductase is excluded since it is not unique to denitrification. Using this definition, a bacterium such as Escherichia coli, which can reduce nitrate and nitrite, is not considered a denitrifier because it reduces nitrite to ammonia and does not have denitrifying type enzymes for the reduction of NO or N2O (Simon 2002). In contrast, a bacterium such as Wolinella succinogenes is considered a denitrifier because it contains Nos, which can be used to support growth under anoxic conditions (Yoshinari 1980). Like E. coli, W. succinogenes can reduce nitrate to ammonia but this does not impact its categorization as a denitrifier (Bokranz et al. 1983). Because of this delimiting, nitrate reductase will not be a focus of this chapter.

Phylogenetic Distribution of Denitrification In the pre-genomic era, establishing a bacterium was a denitrifier entailed testing its ability to grow under O2-limiting conditions with nitrate most frequently provided as terminal oxidant (Payne 1981). As a consequence, nearly all denitrifiers characterized were complete denitrifiers that showed robust growth under denitrifying conditions. This perhaps gave a somewhat inaccurate perception that denitrifiers would all grow well under anoxic conditions and produce N2. However, with genome analysis supplanting phenotypic assignment as the principal means of identifying denitrifiers, both complete and partial denitrifiers can be identified, allowing a more accurate assessment of the actual phylogenetic distribution of denitrification genes. The diversity uncovered by this analysis and the often patchy gene distribution makes it unrealistic to provide a complete list of all denitrifiers. Therefore, in the discussion below, only representative examples will be described. Much of this analysis was generated by searches of genomes available through the DOE IMG website (http://img.jgi.doe.gov/cgi-bin/ w/main.cgi). Archaea: Several denitrifying Archaea were characterized in the pre-genomic era, so it has been known for some time that

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denitrification occurred in this group. Genome sequencing has revealed additional denitrifiers. With one exception, all the known archaeal denitrifiers are also capable of aerobic respiration. One review on denitrification in Archaea is available (Cabello et al. 2004). Euryarchaeota: A number of halophilic Euryarchaeota are denitrifiers. This includes members of the Haloarcula, Haloferax, and Halomicrobium genera (Inatomi and Hochstein 1996; Mancinelli and Hochstein 1986). Ferroglobus placidus DSM 10642 is another member of this phylum with denitrification genes. This bacterium is particularly unique in that it has been reported to be an obligate anaerobe capable of denitrification (Vorholt et al. 1997). Recent sequencing of this bacterium’s genome has shown this phenotypic characterization is accurate since it lacks genes encoding proteins required for O2 respiration. Despite this, F. placidus does have a respiratory-based physiology since its main mode of energy generation is to use Fe(II) as reductant and either nitrate or thiosulfate as terminal oxidant (Hafenbradl et al. 1996). Analysis of the genome of F. placidus reveals the presence of a nitrate reductase, a qNor and Nos. Unexpectedly, there is no obvious Nir of either the copper- or heme-containing type. Biochemical data suggest the bacterium has nitrite reductase capacity (Vorholt et al. 1997). However, the rates of NO production were slow and initial characterization of this bacterium suggested nitrite was the major product of nitrate reduction (Hafenbradl et al. 1996). It is strange this bacterium, which is a strict anaerobe, would have lost the capacity to reduce nitrite. Crenarchaeota: Members of the genus Pyrobaculum, which are hyperthermophilic Crenarchaeota, have been found to be robust denitrifiers (Volkl et al. 1993). A qNor has been purified from Pyrobaculum aerophilum (de Vries et al. 2003). Genome analysis indicates this bacterium also has a cd1-Nir, which has not been purified (Cabello et al. 2004). The genome also encodes the genes for a nitrate reductase but lacks the genes encoding the catalytic subunit of Nos. However, this bacterium has been shown to produce N2 as a major end product during denitrification, suggesting the presence of a novel N2O reductase (Volkl et al. 1993). It might be predicted then that the extremophilic nature of this bacteria might be the reason for a novel Nos. However, one member of this genus, Pyrobaculum calidifontis JCM 11548, has a nos gene cluster, suggesting hyperthermophilic conditions do not require a novel protein for N2O reduction. Genome sequencing has revealed several Sulfolobus species, which are also Crenarchaeota, are partial denitrifiers. They have extremely truncated denitrification pathways since they only contain qNor. A few other genera within the Crenarchaeota including Vulcanisaeta, Acidilobus, and Caldivirga also contain species that have a qNor as their only obvious nitrogen oxide reductase. The denitrification components of these Archaea have not been characterized extensively. A copper-containing nitrite reductase has been purified from Haloferax denitrificans and was shown to be spectroscopically similar to related eubacterial nitrite reductases (Inatomi and Hochstein 1996). A CuNir as well as a dissimilatory nitrate reductase have been purified from

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Denitrifying Prokaryotes

Haloarcula marismortui (Ichiki et al. 2001; Yoshimatsu et al. 2000). The qNor purified from P. aerophilum has many similarities to orthologs from Eubacterial sources (de Vries et al. 2003). While limited, these studies confirm the nitrogen oxide reductases of the Archaea are orthologous with those found in the Eubacteria. One other notable archaeon with CuNir is the nitrifying archaeon Nitrosopumilus maritimus. This bacterium has two nirK genes, the designation for genes encoding CuNir, but lacks any obvious Nor or Nos. (Walker et al. 2010). The proteins encoded by these genes have high similarity suggesting they are paralogous. The genome of this bacterium is enriched for genes encoding copper-containing proteins perhaps explaining the CuNir duplication. Nitrite has been proposed as an alternate terminal electron acceptor, but it is not clear how cells would cope with the NO generated by this reaction (Walker et al. 2010). Eubacteria: Denitrification genes are widely distributed among the eubacteria, and current evidence indicates they are only found in eubacteria capable of aerobic growth. Most of the characterized denitrifiers in the eubacteria belong to the proteobacteria. Due to the relative paucity of understanding of denitrification among non-proteobacterial denitrifiers, they have been placed into one section.

Non-proteobacterial Denitrifiers Firmicutes: The frequency of denitrification among firmicutes is uncertain. Importantly though, it is relatively uncommon among most of the model organisms used to study this group. In the genus Bacillus, for example, denitrification is not considered important since most strains of Bacillus subtilis are not denitrifiers. However, a recent study of denitrification in a large collection of Bacillus strains suggested that denitrification occurred in nearly half (Verbaendert et al. 2011). No complete denitrifiers are currently found among those Bacillus species whose genomes have been sequenced. This may only be temporary though since some Bacilli are known to be complete denitrifiers. For example, strains of Bacillus azotoformans have been shown to be denitrifiers (Mahne and Tiedje 1995). More recently, a variety of bacilli were tested for gas production under denitrifying conditions and found to be complete denitrifiers (Jones et al. 2011). Genome sequencing has revealed the potential for partial denitrification in some Bacillus species. For example, qNor is present in Bacillus tusciae strain DSM 2912 and some Bacillus licheniformis strains, but these are their only denitrification enzymes. Denitrification is more common among sequenced Geobacillus than Bacillus species. Some species in this genus are complete denitrifiers, including Geobacillus sp. G11MC16 and Geobacillus thermodenitrificans NG80-2. These are among only a few gram-positive bacteria in which the nos gene cluster has been found. Many others strains including Geobacillus kaustophilus HTA426 lack nos. The G. kaustophilus strain is also notable because it lacks a respiratory nitrate reductase while the others have the genes for this enzyme.

Abbreviated denitrification pathways are found in strains of both Lactobacillus and Staphylococcus. Genome sequencing has found strains in both genera have qNor but lack Nir and Nos. qNor distribution among the staphylococci is somewhat limited with a number of the S. aureus subsp. aureus strains having the gene encoding this protein, while it is totally absent in the S. aureus genomes. The region of the genome containing the gene encoding qNor in these staphylococci is syntenic, suggesting it has not been a recent acquisition. qNor function in these bacteria is likely to detoxify NO, although this has not been demonstrated. Many of these same strains also have a membrane-bound dissimilatory nitrate reductase. Nitrate reductase appears to be coupled to a cytoplasmic NADHdependent nitrite reductase which reduces nitrite to ammonia (Schlag et al. 2008). Several strains of Lactobacillus fermentans have a Nor but are the only lactobacilli to have this gene. These Lactobacillus species possess a nitrate reductase but lack any type of nitrite reductase. Some Lactobacillus such as Lactobacillus plantarum WCFS1 and JDM1 have been found to have nitrate reductase but lack Nir or Nor. In WCFS1, nitrate reductase expression is repressed by glucose and its activity requires exogenous heme and ubiquinol (Brooijmans et al. 2009). Nar activity in these strains can be used to support growth. Another interesting Firmicute with the capacity for nitrogen oxide reduction is Symbiobacterium thermophilum. This bacterium is obligately syntrophic with its growth being dependent on the presence of Geobacillus stearothermophilus (Ueda et al. 2004). Genome analysis has shown that S. thermophilum contains a CuNir and a periplasmic nitrate reductase. Unexpectedly, it lacks Nor. It is uncertain if its Geobacillus partner contains an NO reductase since its genome has not been sequenced. Another Symbiobacterium species, Symbiobacterium toebii, has been shown to be able to grow under anoxic conditions by reducing nitrate (Rhee et al. 2002). Under these conditions, nitrite accumulated stoichiometrically, indicating it lacks a nitrite reductase. Other Firmicutes with Nir while lacking Nor can be found in the genus Thermaerobacter. Thermaerobacter subterraneus and marianensis both have a CuNirK but lack any detectable Nor. They also both lack a nitrate reductase. Actinobacteria: The majority of the actinobacteria capable of denitrification have a severely truncated denitrification pathway. A good example of this is provided by several Actinomyces species which only have only a nitrate and nitrite reductase. For example, genome analysis suggests Actinomyces coleocanis DSM 15436 and Actinomyces odontolyticus ATCC 17982 can reduce nitrate to NO but apparently lack the Nor necessary to reduce this toxic compound to nontoxic nitrous oxide. A number of members of the genus Corynebacterium also have truncated chains. Corynebacterium pseudogenitalium ATCC 33035 has a CuNir but appears to lack all the other nitrogen oxide reductases, including nitrate reductase. Several other strains, including Corynebacterium efficiens YS-314, have both a nitrate and nitrite reductase but lack Nor. A few other strains, including Corynebacterium diphtheriae NCTC 13129, have Nar, CuNir, and a qNor, indicating they could reduce nitrate to N2O but lack an obvious nos ortholog.

Denitrifying Prokaryotes

Another member of this group with a truncated denitrification pathway is Jonesia denitrificans. This bacterium’s genome has been sequenced and it has been found to have a membranebound Nar and a CuNir (Pukall et al. 2009). There are no obvious Nor- or Nos-encoding genes. The Bergey’s description of this bacterium states that it is facultatively anaerobic and can reduce nitrate to nitrite. There is no mention of gas from nitrate in the description, consistent with genome predictions. This indicates that despites its name, Jonesia denitrificans is only a very limited denitrifier. It is puzzling that this bacterium can grow as a denitrifier since the product of its denitrification chain, NO, is toxic. Denitrification seems to be an important trait in Propionibacterium acnes since all sequenced strains contain denitrification genes. All of the strains have a membrane-bound nitrate reductase, a unique CuNir and a qNor but lack Nos. As expected, isolates of this strain have been found to obtain a growth benefit from the presence of nitrate and that nitrous oxide is the final product of nitrate reduction (Allison and Macfarlane 1989). Even though the physiology of Propionibacterium is traditionally associated with fermentation, they are aerotolerant and encode genes whose products are associated with O2 respiration, indicating respiration is an important part of their bioenergetic capacity. Sequence of the skin microbiome has found Corynebacterium, Propionibacterium, and Staphylococcus species are present in high numbers (Kong 2011). Since denitrification is common in these genera, it suggests nitrate or other nitrogen oxides are frequently present in this environment. This has been borne out by the finding that both nitrate and nitrite are present in sweat although the source of these compounds is uncertain (Mowbray et al. 2009). Mycobacterium is another medically important genus in the Actinobacteria whose members contain denitrification genes. Strains including Mycobacterium avium 104, Mycobacterium intracellulare ATCC 13950, and Mycobacterium parascrofulaceum ATCC BAA-614 contain a qNor but no other denitrification genes. It is likely this protein is used for mitigating NO toxicity. However, in this context, it is perhaps unexpected that none of the Mycobacterium tuberculosis strains, which will encounter host-produced NO during infection, contain an NO reductase or other denitrification genes (Voskuil et al. 2011). Assorted others: A variety of other non-proteobacteria species have bits and pieces of the denitrification pathway. Various strains in the family Flavobacteriaceae, which are members of the Bacteroidetes phylum, are partial denitrifiers. Some, like Capnocytophaga ochracea strain ATCC 27872, have only one gene, in this case a qNor. In contrast, the related Capnocytophaga gingivalis JCVIHMP016 lacks Nor but has a CuNir and a Nos. Another pattern is exhibited by Capnocytophaga sputigena Capno which has a CuNir and a qNor. These bacteria are additional examples of bacteria associated with human surfaces since all three were isolated from the oral cavity (http://www.homd. org/index.php#). Nonhuman associated members of the family Flavobacteriaceae also contain denitrification genes. Robiginitalea biformata HTCC2501, which was isolated from the Sargasso

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Sea, encodes a Nos but no other denitrification genes. The same is true for Gramella forsetii KT0803, which was isolated from the North Sea. In contrast, Maribacter sp. HTCC2170 which was isolated off the Oregon coast has the complete set of denitrification genes including cNor. This bacterium, along with Marivirga tractuosa DSM 4126, is the only member of the phylum Bacteroidetes that has a cNor. The latter also has a CuNir and a Nos. Flavobacterium johnsoniae UW101, a soil isolate, has CuNir and qNor but lacks Nos. A few cyanobacteria contain denitrification genes. Analysis of the genome sequence of Synechocystis sp. PCC 6803 showed that it encodes a qNor and a transcriptional regulator that may regulate expression of this enzyme (Kaneko et al. 1996). The same is true for Arthrospira sp. PCC 8005. Acaryochloris marina MBIC11017 has two related copies of genes encoding a quinoloxidizing Nor. It lacks all other denitrification genes. It is not obvious why any of these free-living bacteria would require the activity of a NO-reducing enzyme. Several members of the genus Chloroflexus, including Chloroflexus aurantiacus J-10-fl and Chloroflexus sp. Y-400-fl, have a CuNir. Unexpectedly, this is the only denitrification gene they possess. Chthoniobacter flavus Ellin428, a member of the phylum Verrucomicrobia, also only has a CuNir. Opitutus terrae PB90-1, another member of the Verrucomicrobia, has a CuNir and Nos but lacks genes for Nor. Two members of the Leptospira, which is within the phylum Spirochaetes, are denitrifiers. Both strains of Leptospira biflexa Patoc 1, Ames and Paris, are complete denitrifiers. They possess cNor and a CuNir. These are nonpathogenic, saprophytic members of this genus (Picardeau et al. 2008). None of the pathogenic Leptospira have denitrification genes. One of the most interesting denitrifiers isolated recently is Candidatus ‘‘Methylomirabilis oxyfera.’’ This is a novel methylotroph capable of coupling methane oxidation to denitrification (Ettwig et al. 2010). Metagenome analysis has revealed the presence of a cd1-Nir and three copies of genes encoding qNor. The genome lacks the nos gene cluster which was unexpected since a culture highly enriched with this bacterium produced N2 from nitrite, which is the preferred nitrogen oxide substrate. Recent studies suggest N2O is produced by a novel mechanism in which two molecules of NO are dismutated to produce N2 and O2 with the liberated O2 being used for methane oxidation (Wu et al. 2011). Proteobacteria: The majority of denitrifiers are found in the phylum Proteobacteria. The proteobacteria have been subdivided into five classes: a, b, g, d, and e. Denitrifiers have been found in all of these subdivisions and so each will be discussed separately. d Proteobacteria: While this subdivision contains a number of strict anaerobes some members do possess denitrification genes. A number of these are members of the genus Geobacter, including Geobacter metallireducens strain GS-15, Geobacter sp. M18, and Geobacter sp. FRC-32 which have only the qNor (> Fig. 10.3). G. metallireducens strain GS-15 can grow anaerobically with nitrate but it reduces it to ammonia (Lovley and Phillips 1988). Geobacter bemidjiensis strain Bem

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. Fig. 10.3 Phylogenetic tree derived from Nor sequences from the D proteobacteria. Strains with more than one Nor are indicated by colored boxes, with each Nor from a single strain having the same color. qNor and cNor variants are indicated by the bracketed regions

and Geobacter sp. strain M21, also have qNor. Unexpectedly, these two strains also have cNor as well (> Fig. 10.3). No strain has the cNor alone. The qNor in G. metallireducens strain GS-15 is more closely related to the qNor in Desulfovibrio sp. FW1012B, another d proteobacterium, than other Geobacter strains. Several species in the genus Anaeromyxobacter, such as Anaeromyxobacter sp. Fw109-5, have qNor and Nos but lack nitrite reductase. The Fw109-5 strain also contains a cNor but is the only member of this genus to do so. Anaeromyxobacter sp. strain K is noteworthy because it has three copies of genes encoding qNor. Identity between the three copies is low, about 40 %, making it difficult to determine if the genes are paralogous. These bacteria also can reduce nitrate to ammonia (Sanford et al. 2002). Another interesting member of the d subdivision with a partial denitrifying pathway is Bdellovibrio bacteriovorus HD100, which has a CuNir and a cNor but no Nos (Rendulic

et al. 2004) (> Fig. 10.3). The nirK gene product forms a clade with Nir from g proteobacteria including Ralstonia and Burkholderia. The nor gene cluster of this bacterium is most closely related to putative orthologs from Leptospira, which are members of the phylum Spirochaetes. Since Bdellovibrio are bacteriovorus, it is possible these genes were acquired via some type of horizontal gene transfer (HGT). The G+C content of the Bdellovibrio genome is about 51 %. If HGT was the source of these genes in Bdellovibrio, this might be detected by anomalous G+C content. The bacteria with the most closely related CuNir have a higher G+C than Bdellovibrio while the Leptospira species have a lower G+C content than Bdellovibrio. However, the genes encoding the CuNir and cNor in Bdellovibrio have near backbone average G+C content, so their exact source remains unclear. One other interesting member of the d subdivision is Syntrophobacter fumaroxidans MPOBT. This bacterium was

Denitrifying Prokaryotes

isolated from anaerobic sludge and is a propionate oxidizer in the presence of hydrogen- and formate-utilizing bacteria (Harmsen et al. 1998). It is described as a strict anaerobe but its genome revealed the presence of a bd-type oxidase that could be used for O2 respiration as well as the presence of a qNor but no other denitrification genes. e Proteobacteria: A number of strains of Campylobacter, which, in general, are important human pathogens, have truncated denitrification chains. For example, Campylobacter concisus (strain 13826) and Campylobacter curvus (strain 525.92) both have a quinol-oxidizing Nor and Nos. Campylobacter rectus RM3267 and Campylobacter fetus subsp. fetus strain 82-40 lack the Nor but retain Nos. Other members of the order Campylobacterales such as Wolinella succinogenes also only have a Nos present in the genome. This bacterium along with several of the Campylobacter have been shown to be able to grow using N2O as a terminal electron acceptor (Payne et al. 1982; Yoshinari 1980). It is interesting though that none of these bacteria have a NO-producing Nir, requiring that either the NO or N2O needed for denitrification be produced by an exogenous source. Sulfurimonas (Thiomicrospira) denitrificans ATCC 33889, another member of the order Campylobacterales, is a complete denitrifier with a heme type nitrite reductase and a cNor. It also has two copies of the gene encoding the substrate-binding subunit of Nos in its genome but only one set of the accessory genes required for assembly of the copper-containing cofactors. This bacterium was isolated from a hydrothermal vent (Sievert et al. 2008). Other e proteobacterial denitrifiers have been isolated from hydrothermal vents such as, Nitratiruptor sp. SB155-2 which is a complete denitrifier with a cd1-Nir and a cNor (Nakagawa et al. 2007). Nitratifractor salsuginis DSM 16511 was isolated from the same hydrothermal vent system as the Nitratiruptor sp. and also has a cd1-Nir, cNor, and Nos (Nakagawa et al. 2005). These latter two bacteria are not members of the Sulfurimonas. a Proteobacteria: Denitrification is a very common trait among the a subdivision of the proteobacteria. Probably the best studied denitrifier, Paracoccus denitrificans, is a member of this group (Baker et al. 1998). This bacterium is a complete denitrifier with a cd1-Nir and a cNor. While complete denitrification is common among members of this subdivision, there is also significant variation in pathway length, reflecting the varied roles of the N-oxide reductases and the evolutionary histories of the various strains. A good example of this is provided by the photosynthetic a-proteobacterium Rhodobacter sphaeroides. Rhodobacter sphaeroides 2.4.3 has the genes for all four nitrogen oxide reductases; however, a transposase IS4 family protein has disrupted the nos operon, resulting in N2O being the terminal pathway product (Hartsock and Shapleigh 2010) (and unpublished). R. sphaeroides KD131 has the genes for CuNir, cNor, and Nos but lacks a dissimilatory nitrate reductase (Lim et al. 2009). The type strain, R. sphaeroides 2.4.1, only has Nap and cNor (Mackenzie et al. 2001). The remaining sequenced strain, 2.4.9, only has cNor. So the only N-oxide reductase structural genes they have in common are the ones encoding

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Nor, which are located in the same location on chromosome I in all the strains. The gene for CuNir is also located in a syntenic region of chromosome I in both 2.4.3 and KD131. Both 2.4.1 and 2.4.9 appear to have had nirK but have lost it by deletion. The nitrate reductase is on a plasmid in 2.4.1 that is not present in 2.4.9, which otherwise is highly similar to the type strain (Choudhary et al. 2007). The nitrate reductase common to 2.4.1 and 2.4.3 is also on a plasmid but not in a syntenic region. The 2.4.3 strain encodes a second nitrate reductase that is located on chromosome II. This second nitrate reductase is of a type more commonly found in members of the gproteobacteria, suggesting acquisition via HGT. The nos gene cluster is found on chromosome II in 2.4.3 but on a plasmid in KD131 and there is no conservation of flanking genes. All the denitrification genes in these strains have G+C content and codon usage bias that is indistinguishable from backbone norms. As mentioned above, the ability to grow under denitrifying conditions was the main test for determining if a bacterium was a denitrifier in the pre-genomic era. It can be seen from the example with R. sphaeroides that only the 2.4.3 strain would likely show up in a phenotypic survey. However, even this is somewhat in doubt since 2.4.3 grows quite poorly as a denitrifier (Michalski and Nicholas 1988). A similar result was observed with Agrobacterium tumefaciens, which has genes for Nap, CuNir, and a cNor but lacks genes for Nos. This bacterium is frequently referred to as aerobic because its denitrification growth rate is quite slow, particularly compared to robust denitrifiers such as P. denitrificans. This poor growth trait is probably common among natural isolates and would lead to an underestimation of denitrifiers in the environment if culture techniques were the primary method of enumeration. Other interesting denitrifying members of this subdivision include a number of strains which have Nir but lack Nor and Nos. For example, there are currently six members of the family Phyllobacteriaceae with sequenced genomes. Two of these, Mesorhizobium opportunistum WSM2075 and Chelativorans (formerly Mesorhizobium) sp. BNC1, have a denitrification pathway consisting only of a CuNir. The latter strain actually has two copies of the nitrite reductase gene. A third member of this family, Parvibaculum lavamentivorans DS-1, has a CuNir and a qNor. The CuNir of the latter does not cluster with the CuNir of the other two Phyllobacteriaceae. Instead, it clusters with a group of orthologs found in bacteria such as Azospirillum and Hyphomicrobium in a larger group that also includes Polaromonas and Nitrospira, which are b-proteobacteria (> Fig. 10.4). The qNor in P. lavamentivorans DS-1 is also not closely related to proteins from other a proteobacteria but is more closely related to orthologs from Rhodanobacter. These data suggest the denitrification genes in these Phyllobacteriaceae were acquired by HGT, but as with the other examples, examination of DNA sequence does not provide evidence in support of this conclusion. The Nir-only pathway variation is not restricted to the Phyllobacteriaceae since Octadecabacter antarcticus 307, a member of the family Rhodobacteraceae, also only contains a pathway consisting of only CuNir.

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. Fig. 10.4 Phylogenetic tree derived from CuNir sequences from a, b, and g proteobacteria and a few additional non-proteobacteria. a proteobacteria are highlighted in blue, b proteobacteria in green, and g proteobacteria in pink. The strains highlighted in marigold are non-proteobacteria

Some members of this subdivision also contain multiple Nir-encoding genes which seem unlikely to be a result of gene duplication but reflect independent origins. For example, Afipia sp. 1NLS2 has three copies. One of these is significantly longer than the other two due to presence of an additional cytochrome c–binding domain. The other two proteins are dissimilar from each other, with one grouping with CuNir from Nitrosomonas and Nitrobacter while the other clusters with orthologs from members of the genus Rhodopseudomonas (> Fig. 10.4). The Afipia strain is a complete denitrifier and it is

uncertain if the three nirK are functionally redundant or serve distinct physiological purposes. Oligotropha carboxidovorans strain Om5 also has two distinct nirK genes with one having a c-type cytochrome domain while the other does not. This bacterium lacks Nor but has Nos. The region of the genome containing the nos gene cluster as well as the genes encoding Nar is not present in the nearly identical Om4 strain (Volland et al. 2011). Evidence suggests Om5 is the ancestor and the genes have been lost from Om4. Neither nirK is located in this unconserved region.

Denitrifying Prokaryotes

The most heavily sequenced a-proteobacterial genus with denitrifying members is Brucella. Brucella species are zoonotic pathogens that are no longer free-living (Roop et al. 2009). Many strains from a variety of mammalian hosts have been sequenced. Nearly all are complete denitrifiers. This is not too surprising since this genus is in the order Rhizobiales, which includes many other denitrifiers. Since Brucella are intracellular pathogens, it is likely they are undergoing genome reduction (Wattam et al. 2009). Surprisingly, there is not much evidence of an enhanced rate of loss of denitrification genes. All sequenced strains seem to have an intact cNor-encoding gene cluster. Only one sequenced strain, B. neotomae, lacks nirK and has been shown to lack Nir activity (Baek et al. 2004). The presence of nos is more variable. Most strains have an intact gene cluster but in a number there has been an accumulation of stop codons. For example, in B. melitensis ATCC 23457, the nosZ gene, which encodes the catalytic subunit, contains a single stop codon at about the middle of the orf. All of the other nos subunits, required for assembly of the copper-containing prosthetic groups, remain intact. However, the inactivation of nos also occurs in free-living denitrifiers. For example, two strains of Ochrobactrum, free-living close relatives of Brucella, have been sequenced. One, Ochrobactrum anthropi ATCC 49188, is a complete denitrifier, while the other, Ochrobactrum intermedium LMG 3301, lacks the nos cluster (Chain et al. 2011). The O. anthropi strain is another example of an a-proteobacterium with two nirK genes. One has the nirV gene that has been shown to form an operon with nirK in many a-proteobacteria (Jain and Shapleigh 2001) and is in a region of the genome with other denitrification genes. The second copy lacks nirV and is not proximal to other denitrification genes, suggesting it may not be required for denitrification. The two genes are not closely related (> Fig. 10.4). In general, most a-proteobacterial denitrifiers possess cNor. However, a few bacteria such as Sphingomonas wittichii RW1 have qNor. Another a-proteobacteria with a qNor is Pseudovibrio sp. JE062, which is a complete denitrifier, and its Nor is related to the one found in S. wittichii. Interestingly, Pseudovibrio is another bacterium with both qNor and cNor. The genes encoding the latter are found in a cluster with other denitrification genes, suggesting it may be used during denitrification growth. The function of the qNor is uncertain. The occurrence of Nir is not so biased, with both CuNir and cd1Nir denitrifiers being common among a-proteobacteria. Some closely related denitrifiers, such as P. denitrificans and R. sphaeroides, both members of the family Rhodobacteraceae, have different types of nitrite reductases, with the former having a cd1-Nir and the latter a CuNir. Another interesting group of a-proteobacteria with denitrification genes are nitrifiers of the genus Nitrobacter. The three sequenced representatives of this genus all contain a CuNir. These bacteria lack Nor as well as Nos. The Nir in these bacteria is particularly short, only 300 residues versus the 365–370 residue long proteins other Bradyrhizobiaceae, of which Nitrobacter is a member. This shorter version is also found in Sphingomonas wittichii RW1 and some other a-proteobacteria that are not nitrifiers. The function of the nitrite reductase in nitrifiers is

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not clear (Beaumont et al. 2004; Cua and Stein 2011; Kampschreur et al. 2007). b Proteobacteria: The b proteobacterial class also includes many organisms with denitrification genes. The most heavily represented in genome databases are members of the genus Burkholderia. Burkholderia mallei is the causative agent of glanders and is derived from Burkholderia pseudomallei, which can be isolated from tropical soils (Galyov et al. 2010). B. mallei has undergone genome reduction and has lost the ability to survive in the environment. All of the B. mallei strains have CuNir and a qNor but some have lost Nos. All of the sequenced strains of B. pseudomallei appear to be complete denitrifiers. Despite the frequent occurrence of denitrification, there has been very little research on this genus’ denitrification capacity. Genome analysis indicates these bacteria have a number of unique denitrificationrelated traits. Most strains have two CuNir-encoding genes. One of these is adjacent to and may form an operon with a gene encoding a qNor. The other gene does not cluster with other denitrification genes. While each CuNir is highly conserved when aligned against orthologs from other Burkholderia, paralogs have only about 40 % identity (> Fig. 10.4). Interestingly, one of the two forms is lacking a histidine residue known to be a ligand for one of the copper cofactors in CuNir. This indicates this protein would be inactive. This putatively inactive Nir is encoded by the nirK adjacent to nor. However, this is not the only gene encoding a Nor ortholog in Burkholderia. As with nirK, there is a second nor not clustered with other denitrification genes. Also, as with nirK, the nor orthologs are highly conserved but the paralogs less so. Unlike with nirK, both copies of the nor genes possess the His residues required for ligand binding in nearly all the strains. In a few cases, the nor gene adjacent to nirK has lost conserved His required for cofactor binding or has undergone frame shifts. This suggests the clustered nirK-nor region may have been supplanted by other copies of these genes located elsewhere in the genome and is being lost through site-specific mutation and orf inactivation. Two other b proteobacterial genera, Ralstonia and Thauera, have many denitrifying members. These genera are common soil isolates, particularly from environments contaminated with aromatic carbon compounds (Parales et al. 2008). The former is member of the family Burkholderiaceae while the latter is a member of the Rhodocyclaceae. Not all members of the Ralstonia are complete denitrifiers. All sequenced strains have a qNor but some lack the CuNir that is found in others. The nos-encoding gene cluster is also not present in all strains. Cupriavidus necator, which was originally designated Ralstonia eutropha H16, has a cd1-type Nir. This strain also has two copies of the gene encoding qNor. Only one strain of Thauera, Thauera sp. MZ1T, has a completed genome at this time and it is a complete denitrifier. It has a cNor and a cd1-type nitrite reductase. Like many Ralstonia and Thauera strains, Polaromonas naphthalenivorans CJ2 was isolated because of its ability to degrade aromatic pollutants (Jeon et al. 2003). Subsequent genome analysis indicates this bacterium is a denitrifier with a CuNir and a qNor (Yagi et al. 2009). Attempts to grow this bacterium as a denitrifier have failed (Madsen 2011, personal communication).

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Denitrification is a common characteristic of most species in the genus Neisseria, a well-studied member of the b subdivision. All of the Neisseria gonorrhoeae sequenced to date contain genes for CuNir and qNor. The two genes are adjacent but divergently transcribed. All of these strains also contain the nos gene cluster. Even though they have the cluster, they lack Nos activity since the Nos structural gene encodes a truncated protein lacking residues critical for binding one of the copper cofactors (Barth et al. 2009). None of these strains seems to have a nitrate reductase, suggesting that nitrite is the principal nitrogen oxide encountered in their natural environment. Most strains of Neisseria meningitidis also have a nirK and an adjacent qNor-encoding gene (Barth et al. 2009). However, in a couple of strains, these genes are inactivated. In Neisseria meningitidis C, 8013 both nirK and nor are pseudogenes. Interestingly, in Neisseria meningitidis 053442 ST-4821, the nirK has been displaced by a putative transposon. None of the Neisseria meningitidis strains have the entire nos gene cluster. However, there are remnant genes of the nos cluster present, suggesting these strains had nos capacity at some point (Barth et al. 2009). The N. meningitidis strains also lack a nitrate reductase. All of the other Neisseria species sequenced to date also have a nirK and qNor gene, indicating denitrification is more conserved in this genus than in most other genera characterized to any level of detail. However, with the exception of Neisseria mucosa, all lack a nitrate reductase. The b proteobacteria also have several nitrifiers that have denitrification genes. This includes Nitrosospira multiformis ATCC 25196 which has CuNir and a cNor. Another member of the Nitrosomonadaceae family, Nitrosomonas sp. AL212, also has a CuNir and a cNor. g Proteobacteria: This group contains a large number of denitrifiers including the important model denitrifiers Pseudomonas stutzeri and Pseudomonas aeruginosa (Zumft 1997). While denitrification is found in many pseudomonads, many important branches of this genus lack this capacity. For example, all P. aeruginosa strains in the genome database used in this analysis have a cd1-Nir, a cNor, and Nos gene cluster. In contrast, other well-characterized species such as Pseudomonas syringae do not contain any denitrifying strains. Partial denitrifiers can also be found among the pseudomonads. For example, both Pseudomonas entomophila strain L48 and Pseudomonas fluorescens strain Pf-5 have a CuNir but lack Nor- and Nosencoding genes. In general though, partial denitrification seems less prevalent among sequenced pseudomonads. Other g proteobacteria capable of denitrification include Legionella. Most strains sequenced to date contain a qNor but no other denitrification genes. Allochromatium vinosum strain ATCC 17899, a member of the order Chromatiales, also only has a pathway consisting of qNor. Single step pathways are not restricted to qNor-containing bacteria since Methylococcus capsulatus Bath has cNor but no other denitrification genes. While some of the groups discussed above have thermophilic denitrifiers, the g proteobacteria has at least one psychrophilic denitrifier. Pseudoalteromonas haloplanktis TAC125, whose denitrification pathway consists of CuNir and qNor, was isolated from Antarctic coastal waters and is a model bacterium for the

study of adaptation to a psychrophilic environment (Medigue et al. 2005). Genome analysis has shown this bacterium has completely lost the capacity for molybdopterin metabolism, a cofactor essential for nitrate reductase (Medigue et al. 2005). One of the more unexpected truncated denitrification pathways is found in Vibrio orientalis CIP 102891. This strain of V. orientalis has a nos cluster that appears to be capable of producing an active Nos. No other member of the enteric bacteria has any denitrification capacity. Phylogenetic analysis indicates this gene cluster has close relatives in other g proteobacteria including Colwellia and Photobacterium. Another genus in this group which contains a large number of denitrifiers is Shewanella. This is not too surprising given this genus’ respiratory versatility (Fredrickson et al. 2008). Denitrification is not universally conserved in this genus though since Shewanella baltica seem to lack the capacity for denitrification completely, but on the whole, most species seem to have at least some denitrification genes. Some are complete denitrifiers such as the appropriately named Shewanella denitrificans (Brettar et al. 2002). Many are more similar to the Shewanella putrefaciens strains that have a qNor and no other denitrification genes. It is interesting that S. denitrificans has a cNor instead of the qNor found in other Shewanella. The cNor of S. denitrificans is related to the protein found in other members of the g subdivision such as the pseudomonads or Marinobacter. In contrast, the nirK found in S. denitrificans is similar to the nirK found in other Shewanella such as S. amazonensis SB2B, although there is no synteny in the nirK region of their genomes.

Horizontal Gene Transfer Given the seemingly disjointed scattering of denitrification genes described above, it is not unreasonable to suggest that horizontal gene transfer (HGT) has played an important role in the exchange of genes between strains. However, this cannot be demonstrated definitively since, in most cases, denitrification genes do not have anomalous G+C content or codon usage, nor are they located within obvious mobile elements. One exception is provided by the occurrence of a nirK in Tn6061, a transposon (Tn) found in P. aeruginosa (Coyne et al. 2010). This Tn is 26.5 kb in length and is notable because it encodes resistance to a number of antibiotics. The nirK within Tn6061 is not closely related to nirK in other proteobacteria and is rather divergent from most other nirK orfs (> Fig. 10.4). The most closely related proteins are found in Rhodanobacter, which is a g proteobacteria but with a divergent CuNir, and Opitutus terrae strain DSM 11246, which is a member of the phylum Verrucomicrobia. Many of the bacteria with closely related nirK genes also lack a Nor, suggesting the function of the Nir is something other than anaerobic respiration of nitrogen oxides. This may explain why it is present on a Tn that has no other denitrification genes. Recently, HGTof the nir-nor cluster of Thermus thermophilus has been demonstrated (Alvarez et al. 2011). Transfer of the nir-nor region and an adjacent nar region allowed nitrate-dependent anoxic growth of a recipient that had been obligately

Denitrifying Prokaryotes

aerobic. The nir-nor region may be on a megaplasmid that can be transferred between strains via conjugation. Jones et al. (Hallin et al. 2009) have carried out a thorough phylogenetic analysis of Nir, Nor, and Nos. They note a number of examples where the phylogenies based on 16S and denitrification genes are incongruous. Similar analyses done with sequences available at the time this chapter was written have uncovered additional examples of which a few will be mentioned. Others examples include Methylocella silvestris strain BL2, which is a member of the family Rhizobiales of the a proteobacteria. This strain contains a CuNir that does not group with other family members but clusters with bacteria from the b and g subdivisions (> Fig. 10.4). In contrast, the cNor from this bacterium is found within a clade with many other a proteobacteria. A few other examples include the CuNir of Moraxella catarrhalis strain RH4, a g proteobacteria, which clusters with orthologs from a variety of b-proteobacteria; the CuNir from Methylacidiphilum infernorum V4, a member of the Verrucomicrobia phylum, which clusters with CuNir from the proteobacteria; and the qNor of Arcobacter butzleri strain RM4018, an e proteobacterium, which forms a clade with several members of the Aquificae phylum. Other cases of suspected HGT can also sometimes be inferred if a strain within a genus in which denitrification is common contains denitrification genes distinct from other species within its own genus. For example, the nirK of Shewanella woodyi is quite dissimilar to the nirK of other Shewanella but more similar to those of a nitrifier, Nitrosococcus halophilus (> Fig. 10.4). Another example within the same genus is provided by Shewanella denitrificans which contains a cNor while all other strains with NO reductase use a qNor. HGT may also explain why some bacteria have multiple copies of genes which do not seem to be paralogous in origin. For example, Afipia, a member of the genus Bradyrhizobiaceae, has three nirK. One of these is a member of the Group I CuNir that can be identified with the sequence motif TRPHL, while a second has the sequence SSFH(V/I/P) at the same location, indicating it is a member of the Group II family of CuNir (Hallin et al. 2009). The third nirK encodes a protein which has neither motif but is obviously a CuNir. It seems unlikely that evolution of paralogs gave rise to multiple genes representing well-known subsets of CuNir (> Fig. 10.4). Another example of a bacterium with multiple, non-paralogous copies of a denitrification gene is provided by Anaeromyxobacterium strain 109-5. This strain has three copies of a gene that encodes Nor. Two encode qNor and may be paralogous but the third encodes a cNor. Other examples of a single strain with both qNor and cNor were described above. Sphingomonas wittichii strain RW1, an a proteobacterium, has two qNor, which are only about 50 % identical. One of these clusters with qNor from other a proteobacteria while the other is part of a large clade of enzymes from b or g proteobacteria. The occurrence of a denitrification pathway consisting of one enzyme may also indicate acquisition via HGT. For example, the occurrence of qNor in some of the Staphylococcus aureus subsp. aureus strains but not at all in the Staphylococcus aureus strains or most other Staphylococcus species suggests HGT into

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one branch of the Staphylococcus group. A similar explanation likely accounts for the occurrence of nirK in only a few members of the Chloroflexi. CuNir from these bacteria clusters weakly with CuNir from a variety of bacteria from the b subdivision of the proteobacteria. A somewhat related example is provided by Pyrobaculum calidifontis strain JCM 11548, which is the only member of the Crenarchaeota with a nos gene cluster. As discussed above, the other Pyrobaculum species may have a novel Nos, suggesting that the nos cluster has been recently acquired by JCM 11548 (Zumft and Kroneck 2006).

Enzymology of Denitrification As shown in > Fig. 10.1, complete denitrification is a multistep process, requiring four separate enzymes for the reduction of nitrate and three intermediate nitrogen oxides to nitrogen gas. The basic arrangement of the nitrogen oxide reductases is shown in > Figs. 10.5, > 10.6 and > 10.7. The periplasmic location of several of the N-oxide reductases as well as the fact that none of these enzymes are proton pumps makes denitrification a rather inefficient form of respiration (Strohm et al. 2007). A summary description of the nature of these proteins is presented here. Additional information can be obtained from several recent reviews (MacPherson and Murphy 2007; Tavares et al. 2006; Zumft and Kroneck 2006). Nitrate Reductase: Even though nitrate reductase is not a major focus of this chapter, it is useful to present a brief description of the various types of nitrate reductase (> Fig. 10.5). Nitrate reductase catalyzes the two-electron reduction of nitrate to nitrite. Early studies of nitrate reductase activity in cells demonstrated the existence of at least two types

. Fig. 10.5 Electron transfer pathways to the various forms of nitrate reductase. Nap and Nar receive electrons from quinol (QH2) which is oxidized to quinone (QH). Nas receives electrons directly from NAD(P)H

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. Fig. 10.6 Electron transfer pathways from the bc1 complex to Nir in gramnegative bacteria. QH2 and QH indicate reduced quinol and quinone, respectively. The blue sphere represents small electron shuttle proteins such as c-type cytochromes or pseudoazurin

. Fig. 10.7 Electron transfer pathways from the bc1 complex to Nor in gramnegative bacteria. QH2 and QH indicate reduced quinol and quinone, respectively. The blue sphere represents small electron shuttle proteins such as c-type cytochromes or pseudoazurin

(Zumft 1997). One is a soluble assimilatory enzyme, termed Nas, used when nitrate is the nitrogen source. The other is a membrane-associated respiratory enzyme, termed Nar. More recently, a third type of enzyme was found when a dissimilatory nitrate reductase activity was detected in the periplasm of R. sphaeroides IL 106 (Sawada and Satoh 1980). Further work has demonstrated that this periplasmic enzyme, termed Nap, occurs in a wide variety of bacteria, including both denitrifiers and non-denitrifiers such as E. coli and related enteric bacteria (Richardson et al. 2001). Many denitrifiers contain more than one type of nitrate reductase. The membrane-associated Nar enzymes that have been biochemically characterized consist of a three-subunit complex (Richardson et al. 2001; Tavares et al. 2006). Two of the subunits, NarG (a) and NarH (b), form a cytoplasmically located

heterodimer. They are anchored to the membrane by NarI (g) which is sometimes lost during purification. NarG contains molybdenum, bound by the cofactor molybdopterin guanine dinucleotide and a [4Fe-4S] center (Schwarz et al. 2009). NarH contains several [4Fe-4S] centers and a [3Fe-4S] center. The membrane-anchoring subunit NarI typically contains b-type heme. The direct electron donor to the respiratory nitrate reductase is quinol. The electrons from quinol are thought to be transferred through the heme in the membrane-anchoring subunit to the [Fe-S] centers in NarH and then to the molybdenum center in NarG where nitrate reduction occurs. Nap enzymes are typically heterodimers with prosthetic groups similar to those found in the membrane-associated nitrate reductase (Arnoux et al. 2003). In the heterodimeric form, the largest subunit (NapA) binds molybdopterin guanine dinucleotide and a [4Fe-4S] center. The smaller subunit (NapB) binds c-type heme that is required for transfer of electrons to the active site. In many denitrifiers, electrons are transferred from the membrane-associated quinol pool to the Nap complex by a membrane-bound tetra-heme c-type cytochrome, NapC. Some Nap proteins have been found to have two additional subunits known as NapG and NapH (Brondijk et al. 2004). NapH is an integral membrane protein with multiple [Fe-S] centers while NapG is found in the periplasm but also contains [Fe-S] centers. NapG and NapH likely form a dimer involved in transferring electrons from some form of quinol to the reaction center. In some bacteria such as W. succinogenes, the NapC subunit is missing and its function has been suggested to be taken over by NapGH (Kern and Simon 2008). The periplasmic and membrane-bound enzymes can be distinguished in several ways. First, the membrane-bound enzyme is sensitive to micromolar levels of azide, whereas the periplasmic form is not (Bell et al. 1990). Second, the membrane-bound enzyme can reduce chlorate but the periplasmic enzyme is limited to nitrate, a result that led to the development of a useful method to select nitrate reductase mutants (Bell et al. 1990; McEwan et al. 1984). Third, because the active sites of the two enzymes are on different sides of the inner membrane, the differential membrane solubilities of methyl viologen and benzyl viologen can be used to differentiate activities (Carter et al. 1995). Methyl viologen, which is membrane permeant, can be used as an electron source for both enzymes in whole cell assays. Benzyl viologen, which is membrane impermeant, will act as an electron source only for the periplasmic enzyme in whole cell assays. By comparing the nitrate reductase activity determined with each viologen, the relative levels of activity of each form of nitrate reductase can be estimated. While the function of the respiratory nitrate reductase in denitrification is almost always associated with respiration, the physiological role of the periplasmic enzyme is more variable. If Nap is the sole nitrate reductase in denitrifiers that reduce nitrate to N2O or N2, the enzyme is used for denitrification even though it cannot directly contribute to formation of a proton motive force. However, in denitrifiers with both Nar and Nap, the function of the latter is often not respiration but instead redox homeostasis. This explains why the Nap enzyme is often

Denitrifying Prokaryotes

expressed under oxic conditions and is consistent with the observation that the Nap of Paracoccus pantotrophus is expressed at a higher level in medium containing electron-rich substrates than in medium with more oxidized substrates (Gavira et al. 2002; Sears et al. 2000; Tabata et al. 2005). Nitrite Reductase: Nitrite reductase catalyzes the oneelectron reduction of nitrite to NO, the step that differentiates denitrification from other forms of nitrate metabolism (> Fig. 10.6). There are two types of nitrite reductases that are not structurally related and contain different prosthetic groups (Zumft 1997). One contains copper as a redox active metal (CuNir), and the other utilizes heme-bound iron (cd1). They are both located in the periplasm and appear to be functionally redundant (Glockner et al. 1993). In gram-negative bacteria, both types of Nir receive electrons from the bc1 complex via small, mobile electron shuttles (Hartsock and Shapleigh 2010; van Spanning et al. 2007). Either Nir can be found within species within the same genus but have never been found to occur together in a single bacterium. While there are ecophysiological patterns seen in the phylogenetic distribution of the two enzymes in most cases, the reasons underlying the distribution within a genus or higher taxonomic grouping are not obvious. CuNir has been studied extensively, and much is known about its structure and the nature of the copper centers. Enzymes from several different denitrifiers have been crystallized under different conditions and their high-resolution structures determined (Godden et al. 1991; Jacobson et al. 2005; Nojiri et al. 2009a; Tocheva et al. 2004). These studies have revealed that the enzyme is a homotrimer with each monomer containing two copper atoms. One copper is bound by Cys, Met, and two His residues and is referred to as a type-1 copper center. In multi-copper enzymes, including CuNir, the type-1 copper is involved in electron transfer to the active site. The other copper atom in CuNir is bound by three His residues making it a type-2 copper center. Type-2 centers are found in many multi-copper enzymes and are frequently sites of substrate binding. In CuNir, the type-2 center has been shown to be the site of nitrite binding (Tocheva et al. 2004). While there is a common functional design shared by all CuNir, there is significant variation in the primary structure of the CuNir family. The first recognition of this diversity was from studies of Neisseria gonorrhoeae which reported that a highly expressed protein known as AniA was in fact a CuNir (Mellies et al. 1997). This was somewhat unexpected since this enzyme had low sequence identity, 30 %, with previously characterized CuNir. Determination of a high-resolution structure of the AniA from Neisseria gonorrhoeae showed that despite the limited sequence conservation, the overall structure was similar to previously determined CuNir structures and contained both type-1 and type-2 centers and protein folds to this family of enzymes (Boulanger and Murphy 2002). More recently, additional CuNir variants have been described. The CuNir from Hypomicrobium denitrificans is notable because of the presence of an 15 kDa N-terminal extension. This extension contains a second type-1 copper center (Nojiri et al. 2007). Another variant of this type has been described in

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a number of denitrifiers including members of the genus Ralstonia. However, this variant has an extension at its C-terminus and it contains a sequence motif consistent with the binding of a c-type heme (Ellis et al. 2007). This extension also adds 15 kDa to the protein. An even larger variant has been found in the genome of P. acnes (Nojiri et al. 2009b). This protein is about 900 residues long, in comparison to the 350 residues of the prototypical CuNir from bacteria such as Achromobacter cycloclastes or the 500 residues of the aforementioned extended C- and N-terminus variants. The P. acnes protein contains a classic CuNir type enzyme at its C-terminus, but this is preceded by two additional domains. One of these is a copper-binding domain and the other a predicted transmembrane domain of about 400 residues. The function of the transmembrane domain is uncertain. A number of other Actinobacteria, such as members of the genus Rothia as well as J. denitrificans, have a CuNir with similar domain organization. The cd1-type nitrite reductase is a homodimer that contains a single c-type heme and d1 heme molecule per monomer. The d1 heme is a modified tetrapyrrole ring that is partly reduced and has oxo, methyl, and acrylate side groups (Chang and Wu 1986). The high-resolution structures of the cd1-type enzymes from P. pantotrophus (Fulop et al. 1995) and P. aeruginosa (Nurizzo et al. 1997) have been determined. The protein consists of two domains with the c-type heme located in the smaller domain and the d1-heme, which is also the site of nitrite reduction, located in the larger domain. The reason underlying the usage of the novel d1 heme is uncertain. It has been suggested the d1 heme helps foster a catalytic site with a low affinity for NO and higher affinity for nitrite (Rinaldo et al. 2011). NO has a high affinity for reduced metal centers, so ensuring product release is critical for efficient Nir function. Interestingly, both the CuNir and cd1 enzymes also can reduce O2. Early studies often designated the cd1 enzyme a cytochrome oxidase (Wharton and Gibson 1976). The product of O2 reduction by the cd1 enzyme is water (Lam and Nicholas 1969) although it is not clear how four electrons are passed to the O2 in this process, inasmuch as a one-electron reduction is normally carried out. Nitric Oxide Reductase: In a continuing theme, and as mentioned above, there are multiple forms of Nor (> Fig. 10.7). The first form discovered was cNor. The isolation of the genes encoding cNor in P. stutzeri provided significant insight into the structural organization of the enzyme and confirmed that NO is an obligatory intermediate in denitrification (Zumft et al. 1994). Examination of the deduced primary sequence indicated that cNor is related to the heme-copper family of cytochrome oxidases (HCO) (Saraste and Castresana 1994; van der Oost et al. 1994). Although the overall identity of cNor and HCO members is low, a set of six His residues is conserved in pairwise alignments of subunit I of cytochrome oxidase and the equivalent NorB subunit of Nor. In the HCO family, these residues bind a six-coordinate heme, five-coordinate heme, and copper, the latter two metal centers constituting a binuclear center that is the site of O2 binding and reduction (Iwata et al. 1995).

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In NorB, the equivalent His residues ligate a six-coordinate heme, a five-coordinate heme, and a non-heme iron which replaces the copper at the active site binuclear center. Typically, cNor is purified as a heterodimer with its two subunits, NorB and NorC, being integral membrane proteins. NorC is a c-type cytochrome, proposed to accept electron from the respiratory chain and then transfer them to a b-type heme in NorB. This electron is next passed to the binuclear center which is where NO is bound and reduced to N2O. A recent crystal structure of cNor from P. aeruginosa has confirmed the relatedness of the Nor and HCO families (Hino et al. 2010). One important difference between the two structures though is that the Nor structure lacks residues that form channels or hydrogen bond networks used for proton pumping in some HCO. This is consistent with previous work indicating Nor turnover does not directly lead to formation of a proton motive force (Zumft 1997). While the structure provided significant insight into the organization of Nor, the reaction chemistry by which two molecules of NO are bound and converted to one molecule of N2O remains to be elucidated. A second Nor type was first identified in the bacterium Ralstonia eutropha (Alcaligenes eutrophus H16) (Cramm et al. 1997). This bacterium contains two Nor, one designated NorZ, a product of a gene located on the chromosome, and the other designated NorB, a product of a gene located on a plasmid. The products of these genes have significant identity (>90 %) and are functionally redundant. These enzymes also have significant similarity with the NorB subunit of the cNor, with the exception of an N-terminal extension of about 300 residues. This N-terminal extension binds quinol, which serves as the immediate electron donor for this family of enzymes (Cramm et al. 1999). As with cNor, qNor is related to the HCO family. No high-resolution structure of a qNor is currently available. Genome sequencing has revealed that many denitrifiers utilize qNor. As with the two types of Nir, there is no clear phylogenetic distribution of cNor and qNor but qNor seems to be the preferred Nor among many b and g proteobacteria. A third type of Nor has been found in B. azotoformans and is the only example of a Nor purified from a gram-positive bacterium (Suharti et al. 2001). This enzyme is a dimer like NorCB but lacks c-type heme. Another distinguishing feature is the presence of a CuA-type binuclear copper center. This metal center is also found in HCO and Nos (see below). This enzyme has been designated qCuANor since it will accept electrons from quinol (Suharti and Heering, and S. de Vries. 2004). Unlike qNor though, it will also accept electrons from a membrane-bound c-type cytochrome. Nitrous Oxide Reductase: In general, Nos is located in the periplasm and receives electrons from the bc1 complex via electron shuttles (> Fig. 10.7). Nitrous oxide reductases from several complete denitrifiers have been extensively characterized and several high-resolution structures have been obtained (Brown et al. 2000; Haltia et al. 2003; Paraskevopoulos et al. 2006). Excellent reviews on the genetics and biochemistry of Nos are available (Tavares et al. 2006; Zumft and Kroneck 2006). Purification and characterization of Nos was difficult because its

activity is lost in cell extracts. Nutritional studies had identified copper as an essential nutrient for Nos activity (Matsubara and Zumft 1982) and further work led to the isolation of a soluble copper protein which, under the proper conditions, had Nos activity (Zumft and Matsubara 1982). Nos is homodimeric in most preparations, with four coppers per subunit. Additional studies have demonstrated that when the enzyme is purified anaerobically and assayed using reduced viologens as the electron donor, it has the highest specific activity. The high-resolution structures have defined the nature of the copper-containing sites in Nos (Brown et al. 2000; Haltia et al. 2003; Paraskevopoulos et al. 2006). One of the copper centers has been structurally defined as a CuA site. The CuA center was described originally in the HCO family of proteins (Iwata et al. 1995). CuA is a binuclear site liganded by His, Cys, and Trp residues that is used to transfer electrons from an external donor to the active site, CuZ. CuZ is a tetranuclear site only found in Nos. Histidines are the amino acid ligands to the coppers in the site, but a single sulfide is also an important structural component (Haltia et al. 2003). The high-resolution structure has revealed that the actual number of Cu per dimer is 12 not 8 as suggested from earlier work. The reason for the discrepancy is unclear. It should be noted that crystals were made from protein that had been exposed to O2 which is known to impact function in the enzyme. Therefore, it is uncertain if the metal centers in the published structures are identical to those in the active enzyme. As with the other N-oxide reductases, there is variation among Nos proteins. Campylobacter species and a few other related e proteobacteria contain a C-terminal extension that is a c-type heme-binding domain. W. succinogenes has a Nos modified in this way which is known to be functional since this bacterium will grow with N2O as sole terminal oxidant (Simon et al. 2004; Yoshinari 1980). Unexpectedly though, the genome sequence indicates that the nosZ, which encodes Nos, in W. succinogenes is disrupted by an insertion element. Further studies revealed that some portion of the population carry an intact nosZ gene, likely as a result of the resolving of the insertion element (Simon et al. 2004). Quantitative studies have shown that nos is the least prevalent of the denitrification N-oxide reductases in many environments (Bru et al. 2011; Henry et al. 2006; Keil et al. 2011). This is consistent with genome analyses which find the nos cluster being the least common of all the N-oxide reductases. Genome comparison of closely related bacteria provides frequent examples of nos cluster genes undergoing frame shifts or deletion. For example, a number of Brucella strains show a loss of nosZ or one of the other genes in the nos cluster. Moreover, as mentioned above, Neisseria species express a truncated, inactive Nos (Barth et al. 2009). While Nos seems to be lost at a high rate, it is obvious it is beneficial to some bacteria. For example, almost all of the Burkholderia mallei and pseudomallei strains that have been sequenced have an intact nos gene cluster. Moreover, it is notable that W. succinogenes, Capnocytophaga gingivalis JCVIHMP016, and Gramella forsetii KT0803, isolated from bovine rumen fluid, human oral cavity, and seawater,

Denitrifying Prokaryotes

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respectively, all contain a nos gene cluster but lack Nor, indicating that exogenous N2O is an environmentally relevant terminal electron acceptor.

Regulation of Genes Required for Denitrification This section will focus primarily on the regulation of those genes encoding Nir, Nor, and Nos. In most denitrifiers, the expression of the genes encoding these proteins depends on the presence of nitrogen oxides. Therefore, it is natural to describe the denitrification genes as being part of a stimulon, a term that refers to operons responding together to a particular environmental stimulus (Neidhardt et al. 1990). In general, Nir and Nor are part of a NO stimulon and Nos is part of a N2O stimulon. However, it is important to note that denitrification is a secondary process and so its regulation occurs within organism-specific hierarchies of stimulons and regulons required for aerobic and anaerobic growth. This inevitably will lead to diverse regulation and, in fact, not all of the regulatory strategies described here explain all known regulatory patterns and it is certain that as additional research on regulation of the N-oxide reductases is carried out, additional regulatory patterns will be encountered. Nevertheless, the general patterns described here are consistent with regulatory models developed for many model denitrifiers. One important consideration in the regulation of denitrification is the role of O2 in controlling expression of denitrification genes. While denitrification is traditionally considered as an anaerobic form of respiration, there are many reports in the literature of aerobic denitrification (Kim et al. 2008; Wan et al. 2011; Xie et al. 2003). Studies with model organisms have consistently found that reduction of nitrate or nitrite to gaseous products occurs at low O2 (Shapleigh 2011; Zumft 1997). In particular, it seems that the regulation of nir and nor genes is particularly sensitive to O2 and it is their expression that sets the upper limit of O2 at which denitrification can occur (Bergaust et al. 2011, 2008; Korner and Zumft 1989). None of the reported aerobic denitrifiers have been characterized in sufficient detail to determine if they expresses nir and nor in an O2-independent manner or at O2 levels higher than observed in current studies of model denitrifiers. Until this occurs, aerobic denitrification will remain unverified. When considering aerobic denitrification, it is important to note that the regulation of nitrate reductase is not so tightly controlled by O2. There are examples of Nap-type nitrate reductases that have been demonstrated to be expressed under fully aerobic conditions (Gavira et al. 2002; Hartsock and Shapleigh 2011; Tabata et al. 2005). Another important consideration in evaluating reports of aerobic denitrification is that, with the exception of Nos, none of the N-oxide reductases are inhibited by O2. So, unless care is taken to ensure that cells used in aerobic denitrification experiments have been maintained under aerobic conditions, it is possible that proteome carryover may lead to N-oxide reductase activity that is misattributed to aerobic denitrification.

. Fig. 10.8 Representation of the regulation of the nitrogen oxide reductases in Rhodobacter sphaeroides 2.4.3. This bacterium has two Nap enzymes, designated oNap and rNap, with the former being expressed aerobically and the latter being expressed under microoxic conditions (Hartsock). Lines in the cytoplasm represent genes and transcripts. oNap is green to indicate it is expressed under oxic conditions. Lines from PrrB and NnrR ending in arrows indicate positive regulation while the one ending in a line indicates negative regulation. Transcriptional regulators of rNap are unknown, so the only known controlling factor, lack of O2, is indicated. See text for description of regulatory factors

Nir and Nor Regulation: A number of early studies demonstrated that expression of both Nir and Nor in some model denitrifiers depended on Nir activity (de Boer et al. 1996; Tosques et al. 1997; Ye et al. 1992; Zumft et al. 1994). The observation that it was Nir activity and not nitrite reductase per se that was required for the expression of genes encoding Nir and Nor suggested that a product of nitrite reduction was required for gene expression (> Fig. 10.8). An obvious candidate for the likely effector is NO, a possibility consistent with the observation that addition of NO generators to nitrite reductasedeficient strains results in expression of both Nir and Nor genes (Kwiatkowski and Shapleigh 1996; Van Spanning et al. 1999). Moreover, trapping of NO by hemoglobin decreases expression of nir and nor genes (Kwiatkowski and Shapleigh 1996). The accumulated evidence strongly indicates that it is the production of NO that stimulates expression, but activation by a derivative of NO has not been excluded. The primary regulator of the genes that are part of the denitrification-related NO stimulon has been identified in a number of denitrifiers and in most cases is a member of the FNR/CRP family of transcriptional regulators (Korner et al. 2003). This protein has been variously designated NnrR, NNR, or DNR. An excellent review on this family of proteins has

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recently been published (Korner et al. 2003). Recent work has suggested this protein binds heme, explaining its ability to bind NO with high affinity (Castiglione et al. 2009; Giardina et al. 2008). Computational studies as well as mutagenesis studies have determined the binding sites targeted by some members of this family (Rodionov et al. 2005; Hartsock and Shapleigh 2010). Recent work suggests the regulon controlled by the NNR/DNR family is small. In R. sphaeroides, NnrR likely only has eight binding sites which affect a total of 13 genes (Hartsock and Shapleigh 2010). The regulon of the primary NO-dependent regulator controlling expression of Nir and Nor expression in Neisseria meningitidis is similar in size (Heurlier et al. 2008). This regulator, named NsrR, is not a member of the FNR/CRP family. It acts as a repressor while the members of the DNR/NNR family are activators. Not all denitrifiers control nir expression via an NO stimulon. In some Rhizobium strains, members of the NO-responsive NNR family have been found to be important for expression of the nor genes (Mesa et al. 2002). nirK expression appears to be outside the NNR regulon and NO stimulon because it is regulated by a gene termed FixK2, a member of the FixK family shown to be critical for regulation of many processes related to nodulation in these bacteria (Torres et al. 2011). FixK2 itself is under control of FixJ, a global regulator of low O2 gene expression in rhizobia. Some denitrifiers use additional layers of regulation to link expression of nir and nor to O2 availability. Agrobacterium tumefaciens uses the two component sensor regulator pair ActRS to regulate nirK expression (Baek et al. 2008) (> Fig. 10.8). These regulators are a member of the Reg/Prr family of proteins used by many photosynthetic bacteria to regulate expression of genes whose products are required for photosynthesis. R. sphaeroides 2.4.3 has been shown to use a similar regulatory strategy (Laratta et al. 2002). Members of the Reg/Prr family of regulatory proteins are apparently responsive to changes in the flow of electrons through the electron transport chain (Kim et al. 2007; Swem et al. 2001). As O2 decreases, electron flow through the chain decreases, leading to phosphorylation of the response regulators which are necessary for nirK expression. nor expression is not directly regulated by Reg/Prr but since nirK is, this places nor under indirect control of Reg/Prr since nor is part of the NO stimulon (Baek et al. 2008). Nos Regulation: nos expression is likely controlled by a number of factors, leading to diverse control mechanisms. Evidence for a N2O stimulon was initially provided by the observation that growth on N2O stimulates Nos expression, modestly stimulates expression of the nitrate reductase, and does not stimulate expression of Nir or Nor in P. stutzeri (Korner and Zumft 1989). Additional support for NO-independent expression of nos has been provided by studies of P. denitrificans, where it was shown that expression of nos preceded that of nirS and nor (Bergaust et al. 2011). Use of N2O as an effector is not universal though since P. aeruginosa grown on N2O alone will not express Nos. Nos is expressed when the cells are grown on

nitrate and nos is likely within the NO stimulon (Arai et al. 2003). In P. stutzeri, while N2O alone can activate expression of nos, growth on nitrate and subsequent production of NO leads to even higher expression of the nos gene cluster (Vollack and Zumft 2001). This suggests nos is within both the NO and N2O stimulons in this bacterium, which is probably the case in many complete denitrifiers. Little is known about specific regulators required for nos expression outside of the NNR/DNR family. One protein that appears important for nos expression in nearly all denitrifiers is NosR. NosR is an integral membrane protein which contains [Fe-S] clusters as well as flavin (Wunsch and Zumft 2005). The gene for nosR is almost always part of the nos gene cluster. How NosR regulates nos expression and Nos activity is uncertain. Insertional inactivation of nosR leads to a decrease in nosZ transcription (Cuypers et al. 1992; Velasco et al. 2004). However, nosR does not have any obvious DNA-binding motifs and its membrane localization is seemingly inconsistent with it being a DNA-binding protein. NosR may play a role in Nos assembly since removal of regions with conserved motifs led to overexpression of an inactive Nos in P. stutzeri (Wunsch and Zumft 2005). While the vast majority of denitrifiers contain NosR-encoding genes, W. succinogenes lacks any obvious nosR orthologs (Simon et al. 2004). Density-Dependent Regulation: In addition to nitrogen oxide and O2-dependent control, some of the denitrification genes in P. aeruginosa show evidence of density-dependent regulation. Expression of nirS, nor, and nos in this bacterium is regulated by members of the DNR family (Arai et al. 1997, 2003). However, it was observed that inactivation of genes responsible for densitydependent expression led to an increase in the rate of N2 production from nitrate (Toyofuku et al. 2007). As expected, expression of the genes encoding Nir, Nor, and Nos showed decreased expression in the presence of an effector. This indicates that under conditions of high cell density, expression of these genes will be repressed. The regulatory mechanisms used by denitrifiers to control Nir, Nor, and Nos expression are consistent with denitrification being nonessential and secondary to O2 respiration. These regulatory strategies also provide evidence of how cells use regulation to ensure denitrification is used to support other more critical physiological processes. For example, in rhizobia, the Fix system of regulators that controls Nir expression is critical for regulating a variety of genes required for nodule formation and diazotrophy (Mesa et al. 2008). Similarly, photosynthesis is a major mode of energy generation in R. sphaeroides and so it is not unexpected to find the denitrification genes are regulated by global regulators involved in adapting cell physiology for photosynthetic growth, also a low O2 process (Kaplan et al. 2005). Density-dependent regulation in P. aeruginosa provides another example. This bacterium uses biofilms as a key means of survival and growth, and biofilm formation is controlled in part by density-dependent regulatory systems (Harmsen et al. 2011). Biofilms often have low O2 microenvironments due to O2 gradients established by respiration. Therefore, it is not surprising to find denitrification genes are controlled by the regulators also

Denitrifying Prokaryotes

required for the formation of biofilms. NO has been shown to convert sessile cells to motile cells which may explain why denitrification genes are repressed by the density-dependent regulators (Barraud et al. 2006).

Environmental Studies There have been many efforts to characterize denitrifier communities in the environment. As might be expected from the broad phylogenetic distribution of denitrification genes, nearly all environments have denitrification genes (Bru et al. 2011; Keil et al. 2011; Philippot et al. 2009). Quantitative assessment of the frequency of occurrence of individual genes has found that genes for nitrate reductase occur at the highest frequency, likely representing its use in both denitrification and DNRA. Genes encoding Nir occur at a lower frequency than Nar and Nap genes. Genes encoding Nos occur at the lowest frequency. Nos activity also appears to be inhibited at low pH (Liu et al. 2010; Simek and Cooper 2002; Thomsen et al. 1994). Low pH environments are often found to produce large amounts of N2O under denitrification conditions (Simek and Cooper 2002). Analysis of the distribution patterns of particular gene types has identified some large-scale trends. A thorough analysis of the distribution of nirK and nirS, the structural gene for the cd1-Nir, has been carried out by Jones and Hallin (Jones and Hallin 2010). They found that nirS sequences were more enriched in saline environments. However, nearly all of these studies are reliant on the use of oligonucleotide primers that may not work with divergent or novel genes. For example, sequencing of the genome of a denitrifying Rhodanobacter strain showed that standard PCR primers would be unlikely to amplify the nirK from this bacterium (Green et al. 2010). This is significant because members of the genus Rhodanobacter are common denitrifiers in low pH environments. It is well established that low pH has been shown to impact denitrifier populations in general, so it is possible that diversity is being underestimated by the use of standard primer sets (Deiglmayr et al. 2004; Green et al. 2010; van den Heuvel et al. 2010). While denitrification genes may be common in most environments, in soils, there is a growing appreciation of the fact that denitrification is ‘‘spotty.’’ Certain sites are ‘‘hot spots’’ relative to other closely located sites (McClain et al. 2003; Vidon et al. 2010). Even within this hotspot framework, it has become obvious that sites are also subject to ‘‘hot moments.’’ Molecular studies have supported the idea of hot spots (Enwall et al. 2010). The occurrence of hot spots and hot moments makes it difficult to accurately quantitate denitrification rates in soils. While denitrification is common in soil and aqueous environments, it is important to note that human-associated communities have the capacity for denitrification. Human saliva is enriched for nitrate, so it is not surprising that denitrification has been found to occur in human dental plaque (Lundberg and Govoni 2004; Lundberg et al. 2004). A number of partial denitrifiers have also been identified during efforts to sequence the

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human oral microbiome (http://www.homd.org/modules.php? op=modload&name=HOMD&file=index&taxonomy=1). Also, as mentioned above, a number of pathogens within the Neisseria, Brucella, and Pseudomonas genera are complete denitrifiers. Denitrification genes in these bacteria have been found to be pathogenicity determinants (Baek et al. 2004; Haine et al. 2006; Hassett et al. 2002; Laver et al. 2010; Stevanin et al. 2007; Tunbridge et al. 2006).

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11 Dinitrogen-Fixing Prokaryotes Ernesto Ormen˜o-Orrillo1 . Mariangela Hungria2 . Esperanza Martinez-Romero1 1 Genomic Sciences Center, UNAM, Cuernavaca, Mexico 2 Embrapa Soja, Londrina, Brazil

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427

Introduction

Diazotroph Isolation and Conditions for N2 Fixation . . . . 428

Dinitrogen fixation, the biocatalytic conversion of gaseous nitrogen (N2) to ammonium, is an exclusive property of prokaryotes, with only few of them having this capacity. Some are symbionts of eukaryotes from plants and animals to protists. The enzymes responsible for nitrogen fixation are nitrogenases (see section > ‘‘Nitrogenase Structure’’). 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). Biological nitrogen fixation is the main process of incorporation of N2 into the biosphere, contributing to 65 % of the total input of N in Earth, or 96 % of the N input derived from natural processes, thus being considered as the main process to life after photosynthesis. 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 Pre´vost 1994; Zahran 1999). Indeed, agronomically important crops such as soybean (Glycine max), alfalfa (Medicago sativa), pea (Pisum sativum), clover (Trifolium spp.), and bean (Phaseolus vulgaris) obtain substantial amounts of their nitrogen from biological N2 fixation. Worldwide some 44–66 million metric tons of N2 are fixed by agriculturally important legumes annually, with another 3–5 million metric tons fixed by legumes in natural ecosystems, providing nearly half of all the N used in agriculture (Postgate 1982; Smil 1999; Newton 2000; Graham and Vance 2003). One of the long-term goals of N2 fixation research is to select or engineer major cereal crops such as rice (Oryza sativa), maize (Zea mays), and sugarcane (Saccharum spp.), 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 may 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. Symbiotic associations of diazotrophs with eukaryotes as diatoms, corals, and fungi (reviewed in Kneip et al. 2007) cause these organisms to acquire nitrogen-fixing capabilities; the analyses of these symbionts reveal fascinating adaptations and in few cases the evolution of nitrogen-fixing cyanobacteria to become organelles called spheroid bodies in diatoms (Kneip et al. 2008).

Methods for Detecting Nitrogen Fixation . . . . . . . . . . . . . . . . . 429 Distribution of Dinitrogen-Fixing Ability Among Prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 430 Ecology of Dinitrogen-Fixing Prokaryotes . . . . . . . . . . . . . . . . . 432 Free-Living Diazotrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432 Symbiotic Diazotrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432 Nitrogen Fixation in Insects . . . . . . . . . . . . . . . . . . . . . . . . . 432 Other Nitrogen-Fixing Symbioses in Eukaryotes . . . 433 Dinitrogen-Fixing Prokaryotes in the Oceans . . . . . . . . . . . 433 Dinitrogen-Fixing Prokaryotes in Agriculture . . . . . . . . . . . . 434 Inoculants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 434 History of Inoculants and Inoculation . . . . . . . . . . . . . . 434 Determining the Need for Inoculation . . . . . . . . . . . . . . 434 Inoculant Production and Utilization . . . . . . . . . . . . . . . 434 Nitrogen Fixation with Legumes . . . . . . . . . . . . . . . . . . . . . . . . 435 Nitrogen Fixation with Nonlegumes . . . . . . . . . . . . . . . . . . . . 437 Interface Rhizobia-Associative/Endophytic Bacteria . . . . 439 Biochemistry and Physiology of Dinitrogen Fixation . . . . . 439 Nitrogenase Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 440 Nitrogen-Fixation Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 440 Regulation of Nitrogen-Fixation Genes . . . . . . . . . . . . . . . . . 442 Lessons from Genomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 442 Conclusions and Perspectives of Application of Nitrogen-Fixation Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 443 Abstract Dinitrogen fixation is a key process in the N cycle and only carried out by few prokaryotes. Research on dinitrogen fixation includes basic and practical applications: from nif genes to crops, with molecular, genetic, ecological, taxonomic, and agricultural approaches used. Nitrogen fixing rhizobia, which have been used in agriculture for over a 100 years, are excellent research models still leading the knowledge of eukaryotebacteria symbioses. Other less known symbioses of dinitrogen fixing bacteria are reviewed as well as free-living diazotrophs.

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_72, # Springer-Verlag Berlin Heidelberg 2013

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Dinitrogen-Fixing Prokaryotes

Industrial fixation of atmospheric N2 tends to exceed 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 N in soil, marine environments such as oligotrophic oceanic waters (where dissolved fixed-nitrogen content is extremely low; 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 nitrogenpoor 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 papermaking chemicals (Vaisanen et al. 1998). Nevertheless, the presence of a N2-fixing bacterium is not evidence for the occurrence of N2 fixation, as in most cases special conditions as low oxygen or a differentiation process are required for nitrogenase to be expressed. The usefulness of N2-fixing bacteria in bioremediation is also being recognized (Suominen et al. 2000; Prantera et al. 2002). Transformation of contaminating polychlorinated biphenyls was obtained with alfalfa inoculated with Ensifer 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). Dinitrogen fixers are encountered in bacteria and in some groups of archaea. The list of the phyla containing nitrogenfixing bacteria is probably still far from complete but enlarging. Knowledge of N2 fixers is limited, and some not yet identified N2 fixers could be found among the novel bacterial divisions that are mostly as-yet-uncultured (Rappe and 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). The reaction of nitrogen fixation may be represented by the equation N2 + 8H+ + 6e ! 2NH3 + H2 and is coupled to the hydrolysis of 16 equivalents of ATP. The high cost of energy may thus explain why nitrogen fixation was lost in many bacterial lineages when not needed. The possession of N2-fixing genes (see section > ‘‘Distribution of Dinitrogen-Fixing Ability Among Prokaryotes’’) does not confer a selective advantage to bacteria in N-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 (Do¨bereiner 1974). Similarly, very few or no Gluconacetobacter diazotrophicus microorganisms were isolated from heavily fertilized sugarcane plants (Fuentes-Ramı´rez et al. 1993, 1999; Muthukumarasamy et al. 1999), and, perhaps as a result of chemical nitrogen fertilization, the bacterial population had very limited genetic diversity (Caballero-Mellado and Martı´nez-Romero 1994; Caballero-Mellado et al. 1995). Another effect of adding fixed nitrogen (diminished genetic diversity of Rhizobium from bean nodules) was observed when the plants were treated with the recommended level of chemical nitrogen (Caballero-Mellado and Martı´nez-Romero 1999).

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 N-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 N-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. Some 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). 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. Cyanobacteria differentiate into N2-fixing heterocysts that protect nitrogenase from oxygen (Wolk 1996). Light was found

Dinitrogen-Fixing Prokaryotes

to induce circadian rhythms of N2 fixation in the cyanobacterium Synechococcus (Chen et al. 1993). Wheat (Triticum aestivum) 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). Free-living diazotrophs are capable of fixing N2 without a host plant, in general utilize the energy available in the environment, and do not excrete or excrete only part of the NH3 produced. In general, these microorganisms live in the rhizosphere, but others live inside stems, roots, and leaves and therefore are considered as endophytes. In a closer relationship, the symbiotic bacteria use the energy derived from the plant photosynthesis and export the NH3 synthesized to the host plant. The major contribution to the biological N2 fixation process occurs by the mutualistic association with plants belonging to the Leguminosae (=Fabaceae) family. This is one of the largest families of plants, with over 18,000 species classified into around 650 genera, representing approximately a twelfth of all known flowering plants and occupying nearly all terrestrial biomes. The family is divided into three subfamilies: Papilionoideae, Mimosoideae, and Caesalpinioideae. Most grain legumes are classified into the subfamily Papilionoideae, and 97 % of the species within this family are capable of fixing nitrogen, while in the Mimosoideae and Caesalpinioideae, the percentages are lower (Allen and Allen 1981; Polhill and Raven 1981; Giller 2001).

Methods for Detecting Nitrogen Fixation The methods used to quantify plant-associated biological N2 fixation have been recently reviewed (Unkovich et al. 2008) and they include (1) methods related to the analysis of total plant nitrogen, applied to both the nitrogen balance method and the nitrogen difference method; (2) quantification by the ureide-N method in some tropical legumes as soybean and common bean; (3) methods related to 15N isotope, including the 15N-natural abundance, and the 15N isotope dilution; and (4) the acetylene reduction assay. Some of these methods are also used to measure the same process in free-living diazotrophs and bacteria associated with other eucaryotic hosts. N balance method, in which N increases are quantified, is a reliable method if inputs and losses of N are correctly accounted, and the N difference method uses the comparison with a neighbor non-fixing plant species (Unkovich et al. 2008). The ureide method is based on the principle that several legumes (e.g., soybean, bean) transport most of the fixed N2 as ureides (allantoin and allantoic acid), and evaluation of these compounds in comparison to total N (in the xylem sap or in the petiole) will relate to the quantity of N2 fixed (Herridge and Peoples 1990; Unkovich et al. 2008). Several experiments have shown positive correlation between the N-ureide and total plant N or grain yield (Neves and Hungria 1987; Hungria et al. 2006a). The 15N-based techniques have also

11

been thoroughly reviewed (Bergersen 1980; Hardarson and Danso 1993), and one of the advantages is the precise quantification with the use of stable isotopes, therefore not requiring special licenses for the laboratory; however, the use of proper controls in the 15N-natural abundance is very important. The acetylene reduction assay (ARA) has been used for over 30 years to measure nitrogenase activity and as an indicator of N2 fixation (Hardy et al. 1968). The method is based on the principle of alternative substrates of the nitrogenase, one of them acetylene, that is reduced to ethylene that can be easily analyzed by chromatography. Acetylene reduction has been very useful to detect new N2-fixing microorganisms. However, it has been demonstrated that disturbance of soil, roots, and nodules and of the interface root-soil may drastically affect activity of both nodulated legumes and associative N2-fixing systems (Minchin et al. 1986; Hunt and Layzell 1993; Unkovich et al. 2008). Alternatives have been proposed, as the evaluation in flow systems and of the H2 flux, but the assay has not been used for quantification purposes (Unkovich et al. 2008). Novel approaches that use 15N with mass spectrometry (MIMS, multi-isotope imaging mass spectrometry or nano-SIMS) allow the detection of nitrogen fixation in cells in microbial consortia or inside eukaryotic host cells (Lechene et al. 2007; Dekas et al. 2009). These approaches may displace older methods. Using nano-SIMS and fluorescence in situ hybridization nitrogen fixation was found to occur in deep-sea sediments such as in the cold methane seeps that were unknown to be sites for nitrogen fixation (Dekas et al. 2009). 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 anchors to design primers for the synthesis of the nitrogenase genes by means of polymerase chain reaction [PCR] amplification) in the structural nif gene sequences (see section > ‘‘Distribution of Dinitrogen-Fixing Ability Among Prokaryotes’’), namely, nifHDK, found in many microorganisms (Dean and Jacobson 1992; Ueda et al. 1995). However, finding nitrogenase genes is not evidence that nitrogen-fixation activity occurs. 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. Furthermore, with his approach, the description and natural histories of communities of N2-fixing microorganisms may be established more accurately than with traditional microbiological techniques. For example, the nitrogenase reductase (nifH) genes may be amplified by PCR 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). In other cases, homologous or heterologous probes have been used in

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hybridization experiments to detect N2 fixers. Hybridization to macro- and microarrays may reveal the presence and frequency of different N2-fixing prokaryotes (Jenkins et al. 2004; Steward et al. 2004). A microarray with a diversity of oligonucleotide corresponding to nifH genes has been used to detect diazotrophs (Zhang et al. 2007). 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. Since knowledge of the nitrogenase gene diversity has improved (over 27,600, 950, and 400 nifH, nifD, and nifK sequences, respectively, were available in the GenBank database at the time this manuscript was being written), different sets of primers have been designed (Bu¨rgmann et al. 2004) to better amplify nifH genes directly from DNA extracted from various samples. A different method of N2-fixation detection involves the growth of indicator non-N2-fixing organisms in a coculture 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 N-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, N2 fixation could have evolved in methanogenic archaea and subsequently transferred into the bacterial domain (Raymond et al. 2004). Among the domain Archaea, nitrogen fixers (or strains containing nitrogenase genes) are restricted to the phylum Euryarchaeota (> Fig. 11.1). Few out of the over 100 currently identifiable major lineages or phyla within the domain Bacteria have nitrogen-fixing members, namely, Proteobacteria, Cyanobacteria, Chlorobi, Spirochaetes, the Gram-positives (Firmicutes and Actinobacteria), Bacteroidetes, Nitrospirae, and Verrucomicrobia, and probably also some strains within Fusobacteria, Deferribacteres, and Fibrobacteres for which only nitrogenase genes have been detected without evidence of nitrogen fixation (> Fig. 11.1). Dinitrogen-fixing organisms have an advantage over nonfixers in N-deficient but not in N-sufficient 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 nonN2 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 (called nodulation genes, including nod, nol, and noe genes) may be easily lost concomitantly with the symbiotic plasmid (Segovia et al. 1991). Similarly, nonsymbiotic Mesorhizobium strains, that lack a symbiotic island, are found in nature (Sullivan et al. 1996). On the other hand, acquisition of the whole set of symbiotic genes by nonsymbiotic rhizobia in N-depleted soils has been reported in both Mesorhizobium (Sullivan et al. 1995; Sullivan and Ronson 1998) and Ensifer (=Sinorhizobium) (Barcellos et al. 2007). 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 (Klebsiella pneumoniae from clinical specimens) had this capacity (Martı´nez 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-Ramı´rez et al. 2001). Similarly, among aerobic endospore-forming Firmicutes (Grampositive 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 molybdenumindependent 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 root-associated 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

Dinitrogen-Fixing Prokaryotes

11

Cyanobacteria Proteobacteria (δ)

Proteobacteria (γ,α,β)

Proteobacteria (α,β,γ) Firmicutes Proteobacteria (α) Firmicutes

Verrucomicrobia

Nitrospirae

I

Proteobacteria (α)

Aquificae

III

II

Firmicutes

Proteobacteria (α,γ)

Firmicutes Euryarcheota

Euryarcheota

0.05

. Fig. 11.1 Phylogeny of nifH genes encountered in different phyla. nifH clades are indicated with Roman numerals

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, 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 (Bu¨rgmann et al. 2004). Anomalies in the phylogenetic position of Betaproteobacteria have been reported as well (Hurek et al. 1997; Minerdi et al. 2001). The first report on the physiology of N2 fixation within the phylum Verrucomicrobia of which most members remain

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uncultured showed that Methylacidiphilum fumariolicum, a metanotroph, fixes nitrogen at low oxygen concentration and the activity was not inhibited by ammonium. Phylogenies of nifHDK genes showed that the genes cluster with those from the Proteobacteria (Khadem et al. 2010).

Ecology of Dinitrogen-Fixing Prokaryotes Free-Living Diazotrophs The fluctuations of marine diazotroph populations have been analyzed. 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 do not fix nitrogen as efficiently as the nonheterocystous cyanobacteria at the high temperatures of the tropical oceans (Staal et al. 2003). The diversity of marine N2 fixers in benthic marine mats was determined from the sequences of nifH genes. The nifH 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, nutrient-limited, 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).

Symbiotic Diazotrophs The ability to form symbiosis may determine the long-term success of an organism. Nitrogen-fixing symbioses range from rather loose to permanent, characterized by morphological and physiological modifications. Dinitrogen-fixing cyanobacteria form symbioses with diverse hosts such as fungi, bryophytes, cycads, mosses, ferns, and an angiosperm, Gunnera (Bergman et al. 1992). In lichens, sponges, and corals (bipartite or tripartite), cyanobacteria fix nitrogen. Heterocysts, the specialized cyanobacteria cells where nitrogen fixation occurs, are formed by Nostoc punctiforme at the hyphal tips of the fungus Geosiphon pyriformis, a relative of AM fungi. Epiphytic moss in subtropical moist forest seemingly associated with nitrogen-fixing bacteria showed acetylene reduction activity (Han et al. 2010). The identity of the nitrogen-fixing species is unknown and N fixation should be evaluated with other methods.

Research on the symbiosis of cyanobacterial species and hosts, Nostoc-Gunnera, Richelia-Rhizosolenia, and Hemiaulus showed that some bacterial symbionts may be acquired from the environmnent while others are transmitted from one generation to the next; the luggage endosymbiont hypothesis was reviewed (Wouters et al. 2009). Each host diatom harbors a unique endosymbiotic Richelia strain. 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 (Martı´nez-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). 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. 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, 1996). 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 within tRNA genes or flanked by insertion sequences (Cheetham and Katz 1995; Kovach et al. 1996). In Mesorhizobium loti, the symbiotic genes are chromosomally located as in most Mesorhizobium and Bradyrhizobium sp. The range of nodulating bacteria has enlarged. Nodulating Methylobacterium has been reported from Crotalaria nodules (Sy et al. 2001). Strains of Devosia, Ochrobactrum, Aminobacter, Microvirga, Shinella, and Phyllobacterium have also been shown to be root-nodule legume symbionts (Rivas et al. 2002; Valverde et al. 2005; Zurdo-Pinero et al. 2007; Estrella et al. 2008; Lin et al. 2008; Ardley et al. 2011). Surprisingly, some Betaproteobacteria in the genera Burkholderia (Moulin et al. 2001) and Cupriavidus (formerly known as Ralstonia) (Chen et al. 2001) are capable of nodulating legumes. Like Rhizobium and Sinorhizobium spp., some 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).

Nitrogen Fixation in Insects Estimates are that the contribution of insect-borne nitrogenfixing bacteria may be up to 30 kg of N/hectare (ha)/year (Nardi et al. 2002).

Dinitrogen-Fixing Prokaryotes

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 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). Primitive termites have protists in their guts that contain bacterial symbionts, among them, one very aggressive woodeating species Coptotermes formosus contains a Bacteroidete that may not be cultured in the lab, designated Candidatus Azobacteroides pseudotrichonymphae that has both nitrogenfixing and cellulolytic capabilities. These bacteria nifH genes resemble those from termite gut spirochaetes (Hongoh et al. 2008) and not those from Chlorobi that is a closer relative to Bacteroidetes. Previously no nitrogen-fixing bacteria were recognized among Bacteroidetes, so this is a new phylum in the list of phyla with nitrogen-fixing bacteria. Enterobacteria mediate nitrogen fixation in natural populations of the fruit fly Ceratitis capitata. Nitrogen-fixing enterobacteria such as Klebsiella and Enterobacter have been cultured from the fruit fly (Behar et al. 2005). Nitrogen is a limiting factor not only in human agriculture but also in ant agriculture. Symbiotic nitrogen fixation was detected in the fungus gardens of leaf-cutter ants. Nitrogenfixing genes detected were found similar to those from Enterobacter (Pinto-Tomas et al. 2009) or Klebsiella variicola (Rosenblueth et al. 2004).

Other Nitrogen-Fixing Symbioses in Eukaryotes Endosymbionts from marine bivalve species, located in the shipworm gills, are cellulolytic and 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

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(Distel et al. 2002). Imaging with mass spectrometry (MIMS) showed that the bacterial symbionts within the wood-eating marine bivalve Lyrodus pedicellatus fix nitrogen in host cells. Evidence was also presented that the fixed nitrogen was transferred to the host (Lechene et al. 2007). In sea urchins Strongylocentrotus droebachiensis, the nitrogen-fixing activity depended on the diet. Digestive traces exhibited nitrogenase activity (Guerinot et al. 1977). Symbionts from the pinnate diatom Rhopalodia gibba have nif genes that resemble both in sequence and organization those genes from free-living cyanobacteria specially Cyanothece sp. Remarkably, the cyanobacterial symbiont has gone through a genome reduction process and lost photosynthetic pigments among many other genes. The endosymbiont is no longer photosynthetic, and this function may be supplied by the host. R. gibba has not been observed without the bacterial symbiont, and the symbiont does not grow alone in culture media in relation to having a reduced genome (Kneip et al. 2008).

Dinitrogen-Fixing Prokaryotes in the Oceans Dinitrogen fixation in the world oceans was reviewed by Karl et al. (2002). In a deep-sea hydrothermal vent, a methagenomic archaeal was capable of fixing nitrogen at 92 C (Mehta and Maross 2006). This raises by 28 C the upper limit of temperature for nitrogen fixation. Archaeas in consortia with sulfate reducing bacteria are responsible for nitrogen fixation in the deep ocean sediments (Dekas et al. 2009). These ‘‘Fantastic Fixers’’ (Fulweiler 2009) were functionally detected by nano-SIMS that is useful to detect nitrogen fixation in situ. A low representation of nitrogen-fixing gene sequences was found in the metagenomic analysis of the ocean (Johnston et al. 2005) that may be explained by the sampling process with elimination of large size cells that would exclude Cyanobacteria as Trichodesmium and Synechococcus. In coastal sediments, considerable levels of nitrogen fixation have been detected (Fulweiler et al. 2007). Filamentous cyanobacteria including Trichodesmium and Katagnymene were the most abundantly detected in an approach independent of culture using real-time quantitative PCR (Langlois et al. 2008). The distribution of nifH phylotypes was in relation to water temperature. Unicellular uncultured cyanobacteria were diverse and also abundant. The diatom endosymbiont Richelia (Wouters et al. 2009) is considered together with Trichodesmium a major nitrogenfixing bacterium in the ocean (Davis and McGillicuddy 2006). However, there are other marine diazotrophs that have been reported including Proteobacteria (Hewson et al. 2006; Man-Aharonovich et al. 2007) that seem to be widely distributed. High nitrate concentration does not seem to select against all diazotrophs (Karl et al. 2002; Langlois et al. 2008). In the ocean, iron and phosphate (San˜udo-Wilhelmy et al. 2001) seem to limit

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Dinitrogen-Fixing Prokaryotes

nitrogen fixation (Voss et al. 2004). Available phosphate limits N2 fixation not only in agricultural fields but also in the ocean (San˜udo-Wilhelmy et al. 2001).

Dinitrogen-Fixing Prokaryotes in Agriculture Inoculants History of Inoculants and Inoculation The use of legumes in agriculture because of their properties of improving soil fertility dates back from a long time, and one example is given by the Romans that developed the idea of crop rotation with legumes and nonlegume plants to improve soil ‘‘health’’ and quality. However, only in 1813, the interest in chemistry of Sir Humphrey Davy led him to report that the legumes ‘‘seemed to prepare the ground for wheat’’ and speculated that the N came from the atmosphere. In 1838, Boussingault, a French agricultural chemist proved that legumes had higher N levels than cereals and concluded that the atmosphere was the source of this N. Finally, in 1886, two German scientists, Hellriegel and Wilfarth, demonstrated that the ability of legumes to convert N2 from the atmosphere into compounds which could be used by the plant was due to the presence of swellings or nodules on the legume root and to the presence of bacteria within these nodules. The first rhizobia were isolated from nodules soon after, in 1888, by the Dutch microbiologist Martinus Beijerinck, and shown to have the ability to reinfect the legume host and to fix N2 in symbiosis. Two years later, two German scientists, Nobbe and Hiltner, demonstrated the advantages of adding pure bacteria with the seeds at sowing and soon after submitted the first patent describing the use of artificial inoculation. Finally, in 1898, the first commercial inoculant industry, Nitragin, was established in USA (Voelcker 1896; Fred et al. 1932; Smith 1992; Hungria and Campo 2004; Hungria et al. 2005b).

Determining the Need for Inoculation Nowadays, the contribution of the biological N2 fixation process to the N balance in all ecosystems is fully recognized, but it is also known that the inputs are even more critical in the tropics, where the soils have low organic matter and nutrient content and are frequently subject to erosion or inappropriate farm management. In these areas, nutrient depletion may be accentuated by the high cost of fertilizers, especially N sources, the majority of which are imported from developed countries (Hungria and Vargas 2000; Giller 2001; Graham and Vance 2003). For example, Sanchez (2002) estimated an annual average depletion rate of 22 kg of N/ha across 37 countries in Africa. Soil physicochemical exhaustion may lead to depletion of diazotrophic bacteria. Concomitantly, in developing countries, high application of chemical fertilizers also results in constrains of diazotrophic bacteria. Therefore, in both cases, there is need to improve the availability,

quality, and delivery of diazotrophic bacteria to the plants, by means of the inoculation process. Although diazotrophic bacteria are spread in all ecosystems, in several cases, there is need to improve biological N inputs by the addition of inoculants. The practice of inoculation dates from long before the description of the N2 fixation process, with reports of the importance of transferring soil from a field where legumes have been grown to new areas being planted to the same crop to achieve good establishment. One example was the recommendation in England, in the sixteenth and seventeenth centuries, of transferring soil from established pastures of alfalfa (Medicago sativa) to new areas. But it was only after the isolation of the first rhizobia and the establishment of the first inoculant industry that the use of inoculants launched worldwide (Fred et al. 1932; Smith 1992; Hungria and Campo 2004). The main situations where inoculation is needed include the following: (1) when an exotic legume is introduced into a new area where it has not been previously grown and where there is no compatible rhizobia (e.g., soybean in the USA); (2) in soils depleted of diazotrophic bacteria because of different agricultural applications (e.g., high application of chemical N fertilizers); (3) when the soil contains a large population of rhizobia that are compatible, competitive but ineffective with the legume of interest (e.g., several reports with common beans); and (4) when improved strains are needed to sustain highly productive improved cultivars (e.g., modern soybean genotypes). In any of these cases, simple trials can define the need for inoculation and must include a minimum set of three treatments: (1) non-inoculated control without N fertilizer, (2) noninoculated control with N fertilizer, and (3) inoculated without N fertilizer. The comparison of these three treatments will indicate if the indigenous population is effective in fixing N2 or if there is need for inoculation, and it will also indicate if the indigenous or inoculant strains are as effective as the N fertilizer or if strain improvement should take place.

Inoculant Production and Utilization Some aspects related to inoculant production and inoculation will be mentioned in this chapter, and valuable complementary information can be obtained in other reviews (Brockwell and Bottomley 1995; Balatti and Freire 1996; Lupwayi et al. 2000; Stephens and Rask 2000; Catroux et al. 2001; Date 2001; Singleton et al. 2002; Hungria et al. 2005b; Sessitsch et al. 2002). Inoculants consist of a product carrying the desirable diazotrophic strains in an adequate substrate. In general, desirable properties of a good inoculant carrier can be summarized as follows: (1) readily available, uniform in composition, and cheap in price; (2) nontoxic to the bacteria; (3) with high water retaining capacity; (4) easily sterilized; (5) readily corrected to a final pH of 6.5–7.3 or to a pH adequate to the bacteria; and (6) allowing good initial growth of the target organism and the maintenance of high cell numbers during storage (Hungria et al. 2005b). Peat has been the most suitable carrier for inoculant production, since it usually meets these requirements, but different

Dinitrogen-Fixing Prokaryotes

sources of peat vary in their capacity to support rhizobial multiplication and survival (Roughley 1970; Balatti and Freire 1996; Maier and Triplett 1996; Hungria et al. 2005b). Unfortunately, mycobacteria that may represent a risk to human health have been found in peat (De Groote et al. 2006). In the absence of peat, a number of materials fitting the characteristics of a good carrier have been alternatively used with different degrees of success, including vegetable oils, mineral oils, plant materials such as bagasse, silk cocoon waste, sawdust, rice husk, corncob, various clays including vermiculite, perlite mixed with humus, diatomaceous earth, lignite and derivatives, coal, filter mud and charcoal-bentonite, among others (Hungria et al. 2005b). Granular inoculants have also been broadly and successfully used in some countries, as Australia, for soil inoculation, but disadvantages rely on the use of non-sterile substrate and on the lower number of cells per g in comparison to other carriers. For seed inoculation, there are also gels and lyophilized inoculants but representing a low percentage of the market. Worldwide, there is an increase in the number of liquid formulations, due to the facility of sterilization and to the low cost of production (Hungria et al. 2005b). For example, in Brazil, peat inoculants represented 98 % of a market of 12 million doses in 1999, shifting to a market of 60 % of liquid inoculants of a total of 18.7 million doses in 2009. Liquid inoculants must carry cellular protectors, usually proprietary substances to protect the rhizobia. Alginate, xanthans, Fe-EDTA, trehalose, carboxymethyl cellulose (CMC), polyvinylpyrrolidone (PVP), and a variety of other polymers have been listed among the bacteria protectors. Simple procedures of manufacturing liquid inoculants have been developed and may be applied for small-scale procedures. An interesting example was achieved in a project coordinated by Niftal (Singleton et al. 2002), and other procedures are listed by Balatti and Freire (1996). Sterilization is an expensive but key step, because a sterile carrier allows high concentration of cells; furthermore, it reduces the frequency and level of contamination and thus the risk of introducing and disseminating plant, animal, and human pathogens (Lupwayi et al. 2000; Catroux et al. 2001; Hungria et al. 2005b). In countries with high-quality products, sterile products are the general rule or are mandated by legislation, and concentration of cells in inoculation is established in at least 108 cells/g or mL of the product, but with the technology available today, there are several products with concentrations as high as 1010 cells/g or mL. Nevertheless, despite the biotechnological advances achieved in the past decades, unfortunately, a considerable percentage of the inoculants produced in the world are still of relatively poor quality (Brockwell and Bottomley 1995; Lupwayi et al. 2000; Stephens and Rask 2000; Hungria et al. 2005b).

Nitrogen Fixation with Legumes In natural ecosystems, the rates of N2 fixation are sufficient to attend the needs of nodulated plants, but commercial crops require far greater amounts of N. Consequently, to achieve

11

high yields, inoculation with selected strains is often mandatory. Quantification of N2 fixation is a difficult task (Unkovich et al. 2008), but in the studies performed so far with several legumes, the estimates of the contribution of the biological process to the plant N nutrition may be very impressive (> Table 11.1). Exotic grain legumes broadly used as commercial crops need inoculation with compatible rhizobia. One relevant example is that of the soybean, a crop with top priority in South America (Hungria et al. 2005a), USA (Paau 1989), and in many other countries where inoculation is considered a realistic alternative to the increasing use of fertilizers. When soybean was introduced in South America, inoculation was needed to guarantee nodulation and N2 fixation, thus in the 1960s and 1970s, strains were brought from foreigner countries to guarantee grain production in the absence of chemical fertilizers. As the soils were void of natural compatible rhizobia, a new approach was considered in the strain selection program and consisted of isolating and testing hundreds of strains after a period of establishment on the soils. The selection process has shown that it was possible to identify natural variants adapted to the tropical conditions and more efficient and competitive than the parental strains. Concomitantly, the plant breeding program has always considered the symbiotic performance with selected strains used in commercial inoculants. This approach has been very successful, such that high soybean grain yields are obtained in Brazil based exclusively on the N2-fixation process (Hungria and Vargas 2000; Hungria et al. 2005a, 2006a, b). Noteworthy is the economy related to the N2 fixation with the soybean crop in Brazil, estimated at US $6.6 billion/year. The specific requirement of inoculants of most commercial soybean varieties can be attributed to the breeding for grain yield conducted largely in North America in the 1960s and 1970s and then spread to other countries. However, in areas where inoculants are not readily available, soybean growth was limited; promiscuous varieties capable of nodulating with indigenous rhizobia were then developed in a breeding program performed by IITA in Africa (Pulver et al. 1985) and were also identified from early introductions of soybean originated from China (Mpepereki et al. 2000). Nowadays, there are reports that some of the promiscuous genotypes can also respond to inoculation, and efforts have been applied to improve yields of these genotypes. In parallel, more productive non-promiscuous genotypes have been introduced in Africa and will need inoculation. Soybean can also associate with fast-growing strains that were first isolated from soybean nodules and soil from the People’s Republic of China, within the center of origin and diversity of this legume (Keyser et al. 1982); the majority of these strains are classified as Sinorhizobium (=Ensifer) fredii. Some of the fast-growers can be very effective in fixing N2 with the soybean, with successful reports of responses to inoculation in Spain, but the main limitation relies in that S. fredii can only outcompete Bradyrhizobium at high pH, of 7 or 8 (Buendı´aClaverı´a et al. 1994; Hungria et al. 2001). Reinoculation is an important topic to be considered both in the case of exotic plants growing in soils that have been previously inoculated and show an established or naturalized

435

436

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Dinitrogen-Fixing Prokaryotes

. Table 11.1 Estimates of N2 fixation rates with some legumes. In most cases, the values represent the compilation of several studies Species

Common name

N2 fixation rates (kg N/ha)

Reference

Acacia spp.

Acacia

5–50

Sprent and Parsons (2000)

Arachis hypogaea

Groundnut

32–206

Unkovich and Pate (2000)

Cajanus cajan

Pigeon pea

68–88

Giller and Wilson (1991)

Calopogonium muconoides

Calopogonium

64–182

Giller and Wilson (1991)

Centrosema spp.

Centrosema

41–280

Giller and Wilson (1991) Unkovich and Pate (2000)

Cicer arietinum

Chickpea

0–141

Desmodium spp.

Desmodium

25–380

Giller and Wilson (1991)

Gliricida sepium

Gliricida

26–75

Giller and Wilson (1991)

Glycine max

Soybean

0–450

Unkovich and Pate (2000); Hungria et al. (2005a)

Lathyrus sativus

Lathyrus

172–227

Unkovich and Pate (2000)

Lens culinaris

Lentil

5–191

Unkovich and Pate (2000)

Leucaena leucocephala

Leucaena

98–274

Giller and Wilson (1991)

Lupinus albus

Sweet lupin

40–160

Unkovich and Pate (2000)

Lupinus angustifolius

Lupin

19–327

Unkovich and Pate (2000)

Lupinus mutabilis

Bitter lupin

95–527

Unkovich and Pate (2000)

Macroptilium atropurpureum

Siratro

46–167

Giller and Wilson (1991)

Medicago sativa

Alfalfa

45–470

Unkovich and Pate (2000); Russelle and Birr (2004)

Melilotus officinalis

Yellow sweet clover

84

Unkovich and Pate (2000)

Neonotonia wightii

Perennial soybean

126

Giller and Wilson (1991)

Phaseolus vulgaris

Common bean

0–165

Giller and Wilson (1991); Unkovich and Pate (2000)

Pisum sativum

Field pea

4–244

Unkovich and Pate (2000)

Pueraria phaseoloides

Tropical kudzu

115

Giller and Wilson (1991)

Sesbania spp.

Sesbania

7–109

Giller and Wilson (1991)

Stylosanthes spp.

Stylosanthes

4–263

Giller and Wilson (1991)

Trifolium spp.

Clover

67–260

Unkovich and Pate (2000)

Vicia benghalensis

Vetch

125–147

Unkovich and Pate (2000)

Vicia faba

Faba bean

12–330

Unkovich and Pate (2000)

Vigna mango

Black gram

119–140

Giller and Wilson (1991)

Vigna radiata

Green gram

58–107

Giller and Wilson (1991)

Vigna unguiculata

Cowpea

9–201

Giller and Wilson (1991)

Zornia glabra

Zornia

61

Giller and Wilson (1991)

population or in the case of legumes compatible with indigenous strains. Inconsistent responses to inoculant application are frequently attributed to these naturalized/indigenous rhizobia, to the enrichment of populations with the cropping of legumes, or to a combination of both factors (e.g., Sadowsky and Graham 1998; Thies et al. 1991; Thies et al. 1995). There are reports of declines in the response of soybean to inoculation when the numbers of rhizobial cells are as low as 10–20 cells/g of soil (Weaver and Frederick 1974; Singleton and Tavares 1986; Thies et al. 1991). However, with the use of selected strains and highquality inoculants, responses to soybean reinoculation have been reported in soils with 103 cells/g of soil or higher, of up to 106 cells/g of soil. In 74 field trials performed in soils with high population of soybean bradyrhizobia in Argentina, yield was

enhanced by a mean of 14 % in comparison to the non-inoculated treatment, while in 29 field experiments performed in Brazil, reinoculation increased yield by 8 % (Hungria et al. 2006b). Most important, in all of these experiments, there was no response to the application of N fertilizers in any stage of plant growth (Hungria et al. 2006a). Several mutants of soybean cultivars, known as supernodulators and tolerant to nitrate have been obtained (Carroll et al. 1985; Gremaud and Harper 1989; Akao and Kouchi 1992), but they also require inoculation with Bradyrhizobium strains. For indigenous legumes, or legumes that have been grown for a long time in the area, e.g., common bean in Africa and Brazil, responses to inoculation can be erratic. Taken as example common bean, poor nodulation and lack of responses to

Dinitrogen-Fixing Prokaryotes

inoculation in field experiments have been frequently reported worldwide (Graham 1981; Hardarson 1993; Vlassak and Vanderleyden 1997). Explanation for the failure in those trials would rely mainly on a high population of competitive indigenous rhizobia but with low efficiency of nitrogen fixation (Graham 1981; Thies et al. 1991). In addition, common beanrhizobia symbiosis is quite sensitive to environmental stresses, such as high temperatures and soil dryness, leading to low N2-fixation efficiency (Graham 1981; Hungria and Vargas 2000). However, it has been shown that the search—within the natural diversity of indigenous soil population—for effective and competitive strains can be successful. In Brazil, the use of selected strains of Rhizobium tropici has resulted in increases in grain yield of up to 900 kg/ha. Noteworthy is that the responses of common bean to inoculation with elite strains in Brazil were observed in soils with at least 103 cells/g of compatible rhizobia (Hungria et al. 2000b, 2003). A variety of other economically important legumes has important contributions to the N inputs. One important example of legume forage is alfalfa (> Table 11.1), estimated to be one of the five most valuable crops in the world, with a worth value estimated in US 7 billion/year (Howieson et al. 2008). 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 rate of fixation of the tree Acacia dealbata is considered sufficient to replace the estimated loss due to timber harvesting (May and Attiwill 2003), and despite the low number of studies performed so far, some long-term trials had proved that inoculation can improve biomass production of trees (Lal and Khana 1996). The application of green manure can also contribute with high inputs of N. 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 Rhizobium huautlense is also a flood-tolerant symbiosis (Wang and Martı´nez-Romero 2000). Despite not representing a legume-rhizobia symbiosis, the so-called actinorhizal plants that associate with Frankia should also be mentioned, as they are of great value for reforestation; actinorhizal plants belong to eight families (Baker and Mullin 1992; Berry 1994). From the values of N2 fixation reported so far, one may conclude that the contribution of N2 fixation with legumes is fundamental to the global N cycling and to recover and maintain soil fertility. This contribution can be substantially increased by the inoculation with selected strains delivered in high-quality inoculants. Unfortunately, the adoption of legumes in agricultural systems is still low, even in countries that need higher contribution, e.g., estimates in African farming systems are that less than 5 % of area planted to legumes (Giller et al. 2006). However, the prospects for the use of legumes in a variety of new applications are outstanding (Howieson et al. 2008). Concerns should be raised about a higher use of

11

N fertilizers, decreasing the capacity of N2 fixation (van Kessel and Hartley 2000). On the other hand, there are research and extension areas in which in-depth efforts were needed to maximize the contribution of N2 fixation to agriculture: (1) plant improvement (breeding); (2) to alleviate and avoid constrains related to acid soils, soil acidification, soil degradation, desertification, and salinization; (3) to search for new legume genotypes; (4) to allow adequate phosphorus supply and utilization; (5) to stimulate crop rotation; and (6) to improve strains and inoculants (Graham and Vance 2000; Howieson et al. 2008). Unfortunately, despite the great advances achieved in the last decade in the knowledge of diazotrophic bacteria, with an emphasis on the ‘‘omics’’ studies, the field constrains to the biological process have increased, and the efforts toward plant and strain breeding for improved N2 fixation are scarce. Crop production on 33 % of the world’s arable land is limited by phosphorus availability (Sa´nchez 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.

Nitrogen Fixation with Nonlegumes A high impact goal of nitrogen-fixation research has been to extend the process to nonlegumes, and this has promoted the search for diazotrophic bacteria that are associated with agriculturally valuable crops. From a basic research perspective, this has increased our knowledge of their diversity. A critical review of the actual contributions of bacterial N2 fixation to the amount of 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 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 400 kg N/ha (> Table 11.1), while conservative values for bacterial fixation in nonlegumes are 20–30 kg N/ha per year, but higher, substantial values have been also estimated. N2 fixation with nonlegumes has started with research with Azospirillum, dating back to the pioneering work of Dr. Johanna Do¨bereiner (Do¨bereiner and Day 1976; Do¨bereiner et al. 1976). After that, many reports have shown that Azospirillum may promote N2 fixation, growth, and yield of numerous plant species, many of which are of agronomic or ecological importance (e.g., Okon and Labandera-Gonzalez 1994; Bashan and Holguin 1997; Hungria et al. 2010). Inoculants containing Azospirillum have been tested under field conditions with important crops in developing and developed countries, with various degrees of responses. In a survey of 20 years of experiments, Okon and Labandera-Gonzalez (1994) reported that 60–70 % of the experiments showed yield increases due to inoculation with

437

438

11

Dinitrogen-Fixing Prokaryotes

Azospirillum, with statistically significant increases in yield from 5 % to 30 %. In Argentina, in a survey of 273 cases of inoculation of wheat with Azospirillum brasilense, 76 % resulted in a mean yield increase of 256 kg/ha; with maize, 85 % of the cases were successful, resulting in a mean yield increase of 472 kg/ha (Dı´az-Zorita and Fernandez Canigia 2008). In addition, the colonization and contribution of several other rhizospheric and endophytic N2-fixing bacteria have been broadly reported. Sugarcane (Saccharum spp.), rice (Oriza sativa), maize (Zea mays), and wheat (Triticum aestivum) are the Gramineae most extensively studied with regard to N2 fixation, but other crops are being studied as well. Sugarcane has been grown for more than 100 years in some areas of Brazil without N fertilization or with very low N inputs, and removal of the total harvest has not led to decline in yield and soil N levels. This observation suggested that N2 fixation may have been the source for a substantial part of the N used by this crop (Do¨bereiner 1961). From 25 % to 55 % (Urquiaga et al. 1989; Yoneyama et al. 1997) or perhaps as much as 60–80 % (Boddey et al. 1991) of the sugarcane, N could be derived from associative N2 fixation, but skepticism about the occurrence of high levels of N2 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 N uptake by different sugarcane varieties (Urquiaga et al. 1989), declining 15 N enrichment of soil mineral N, carryovers of N from one harvest to the next, and differential effects on control plants during the studies (Urquiaga et al. 1992). The mean estimates of fixed N2 for two sugarcane hybrids grown in concrete tanks ranged from 170 to 210 kg N2 fixed/ha (Urquiaga et al. 1992), and evidences of large differences in N2 fixation among different sugarcane cultivars are compelling. Correction for micronutrient soil deficiencies and high soil moisture seems to be key conditions that promote N2 fixation in sugarcane plants (Urquiaga et al. 1992). 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 Do¨bereiner 1988), Herbaspirillum (Baldani et al. 1986), and Burkholderia (Oliveira et al. 2009). Probably, N2 fixation in sugarcane is performed by a bacterial consortium (Oliveira et al. 2009). 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 wild-type 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 have also 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). G. diazotrophicus (James and Olivares 1997), Herbaspirillum seropedicae, and Herbaspirillum rubrisubalbicans

(Olivares et al. 1996) have been clearly shown to colonize sugarcane plants internally. Colonization by G. diazotrophicus was inhibited by N fertilization (Fuentes-Ramı´rez et al. 1999). Several studies have been carried out on nitrogen balance in lowland rice fields in Thailand (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), among other countries. These studies report a positive balance with estimates of around 16–60 kg of N/ha/crop (App et al. 1986; Ladha et al. 1993). In a N 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 (Boddey et al. 1995; Watanabe et al. 1987). Many different N2-fixing bacteria have been isolated from rice roots. These include Azotobacter, Beijerinckia (Do¨bereiner 1961), Azospirillum (Baldani and Do¨bereiner 1980; Ladha et al. 1982), Pseudomonas (Qui et al. 1981; Barraquio et al. 1982, 1983; 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), sulfur-reducing 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 been long used to fertilize agricultural land throughout the world, most notably rice paddies in Asia. Increases in rice plant growth and increases in N content in the presence of cyanobacteria have been documented by many investigators. Plant promotion may also be related to growthpromoting 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). Alternatively, other bacterial species are being tested to promote rice growth, such as the N2-fixing 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 (Van Traˆn et al. 1996). B. 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

Dinitrogen-Fixing Prokaryotes

(Van Traˆn 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). N2-fixing bacteria associated to maize include Azospirillum, Herbaspirillum, Klebsiella (Chelius and Triplett 2001), B. vietnamiensis (Van Traˆn et al. 1996), Rhizobium etli (GutierrezZamora and Martinez-Romero 2001), Paenibacillus brasilensis (von der Weid et al. 2002), and Klebsiella variicola (Rosenblueth et al. 2004). K. variicola was also found associated with banana plants (Martı´nez et al. 2003). Soil type instead of the maize cultivar determined the structure of a Paenibacillus community in the rhizosphere (Araujo da Silva et al. 2003). In a survey performed by Sumner (1990), thirty-two experiments with cereals were considered to respond positively to inoculation, but there were some negative responses, mostly in wheat. However, positive responses of wheat to inoculation with Azospirillum have been reported, resulting in increase in grain yield of up to 18 % in Brazil (Hungria et al. 2010) and up to 63 % in Mexico (Caballero-Mellado et al. 1992). However, in both studies, the main benefits were attributed to plant-growth promotion properties, and not to N2 fixation. There are an increasing number of reports on the isolation of diazotrophic rhizospheric and endophytic bacteria from several plant species. Sweet potato (Ipomoea batatas) may grow in soils poor on N, and associated N2 fixation has been considered to contribute N to these plants. By a cultivation-independent approach, bacteria similar to Klebsiella, Rhizobium, and Ensifer were inferred to be present as sweet potato endophytes (Reiter et al. 2003). Several diazotrophic genera have also been isolated from important crops as cassava (Manihot esculenta Crantz) (Balota et al. 1997), coffee (Coffea arabica L.) (Jimenez-Salgado et al. 1997), flowers, and fruits, among others. An elucidation of the mechanisms related to the quantitative and qualitative differences of diazotrophic bacteria associated with plants is still missing, and probably, several factors may be related, among them the carbon sources released by the plants (e.g., Christiansen-Weniger et al. 1992).

Interface Rhizobia-Associative/Endophytic Bacteria In nature, several plant species compose the different ecosystems, and in traditional agriculture, plants are grown in crop rotation, succession, or intercropped. It is thus expected that several diazotrophic bacteria may be hosted by different plant species. For over seven centuries, rice rotation with clover (Trifolium spp.) 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 were also encountered inside the rice plant with around 104–106 rhizobia/g (fresh weight) of root.

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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. In nonlegumes (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, 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 N content as compared to non-inoculated 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 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 too low to maintain bacterial growth and N2 fixation (Welbaum et al. 1992; Fuentes-Ramı´rez et al. 1999). In acreage cultivated using S. rostrata-rice rotation, A. caulinodans survives in the soils and rhizosphere of wetland rice (Ladha et al. 1992). A. caulinodans can colonize the rice rhizosphere (specifically around the site of lateral root emergence), can penetrate the root at the site of emergence of lateral roots, and can colonize subepidermally intercellular spaces and dead host cells of the outer rice root cortex (Reddy et al. 1997). Growth stimulation of crops such as wheat and maize inoculated with a R. leguminosarum bv. trifolii strain may not be related to N2 fixation (Ho¨lflich et al. 1995). In Mexico, R. etli was found to colonize maize genotypes (Gutierrez-Zamora and Martinez-Romero 2001). On the other hand, a compilation of studies of co-inoculation of rhizobium with azospirillum has shown benefits in parameters as expression of nodulation genes, nodulation, nitrogenase activity, plant biomass and root growth in chickpea (Cicer arietinum), field pea (Pisum sativum), soybean, common bean, common vetch (Vicia sativa), faba bean (Vicia faba), black medick (Medicago polymorpha), alfalfa (Medicago sativa), winged bean (Psophocarpus tetragonolobus), and white clover (Trifolium repens) (Dardanelli et al. 2008). A better understanding of the complex interactions of associative, endophytic, and symbiotic diazotrophic bacteria with plants should be gained in the next years through approaches as metagenomics.

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

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clarified with the use of 15N. All diazotrophs were thought to use the same two-component nitrogenases (consisting of an iron and a 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 (nitrogenase 2), and the iron-only nitrogenase (nitrogenase 3). 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, N-free media. The V-containing nitrogenase produces around three times more hydrogen than the Mo nitrogenase (Eady 1996). As a result of the reduction of N2 by the nitrogenase, H2 is obligatorily produced, resulting in a lost of energy and electrons of at least 25 %. However, some bacteria possess a second enzyme, hydrogenase, capable of oxidizing the H2, producing ATP, and recovering part of the energy lost in the process (Evans et al. 1985, 1987). 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 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, 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). Some endophytes have been found to release (excrete) riboflavin during N2 fixation (Phillips et al. 1999). Elevated CO2 levels provided to legumes were found to stimulate N2 fixation indicating that N2 fixation was limited by the availability of photosynthate (Hardy and Havelka 1973; Zanetti et al. 1996).

Nitrogenase Structure The classical nitrogenase is a complex, two-component metalloprotein composed of an iron (Fe) protein and a molybdenum-iron (MoFe) protein. 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, and it is the active site of substrate reduction (Hoover et al. 1989). All substrate reduction reactions catalyzed by nitrogenase require the sequential association and dissociation of the two nitrogenase components. 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, anfencoded proteins (AnfDGK; Gollan et al. 1993; Pau et al. 1993). The nifDK genes of A. 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 Nitrogenase genes seem to have gone through a long period of divergence and genes from the most conserved of the nitrogenase genes, nifH fall into different families (http://www.es.ucsc. edu/~wwwzehr/research/database/). 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. 11.2). The products of at least six N2 fixation (nif) genes are required for the synthesis of the iron-molybdenum 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 formation 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),

Dinitrogen-Fixing Prokaryotes

11

Sinorhizobium meliloti nifN

Alphaproteobacteria

H

D

K

E

fixA B

C

X nifA

B

Bradyrhizobium japomnicum nifN K

E

N X

S

B

H

fixR

nifA

fixA

Azorhizobium caulinodans nifA

HI

D

K

E

H2 W fixA B

C X

nifB

Klebsiella Gammaproteobacteria nifQ B

A

L F MZW V S U X N

E Y K

D

H

J

Anabaena vegetative cell nifW X N E

K D’

D

HU S

B

Cyanobacteria Anabaena heterocyst nifW X N E

K

D H U S

B

Methanococcus maripaludis Archaea

nifH

nifD

nifK

nifE nifN nifX

nifI1 = glnB1 nifI2 = glnB2 Chlorobium tepidum Green sulfur (Chlorobi)

nifH

nifD

nifK

glnB glnB

. Fig. 11.2 Arrangements of nif genes in dinitrogen-fixing prokaryotes. The nif gene organization in Methanococcus maripaludis is from Kessler et al. (2001)

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. 11.2). 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 low

substrate specificity might have occurred before the last common ancestor of all living organisms emerged (Fani et al. 1999). The repeated sequences clustered around the nif region of the B. 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).

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Alternative nitrogenase genes, anfH, anfD and anfG (Mo-independent) are found in the termite gut diazotrophs. The sequences of these 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).

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. Cyanobacteria and Archaea have 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). Different regulatory elements 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 very complicated regulatory cascades (Dixon 1998; Nordlund 2000; Forchhammer 2003; Zhang et al. 2003). Very diverse modes of regulation of nif genes have been described that vary between species (D’hooghe et al. 1995; Girard et al. 2000). Detailed studies have been carried out in K. pneumoniae, A. vinelandii, A. brasilense, R. capsulatus, Rhodospirillum rubrum, Sinorhizobium meliloti, or B. japonicum. The most common nitrogenases studied are inactivated by oxygen, and accordingly, the expression of nif genes is negatively regulated by high oxygen concentrations. Different oxygen protection mechanisms have been described (reviewed by Vance 1998). 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), B. 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). 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. 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). 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 (Ensifer, Rhizobium, Azorhizobium and Bradyrhizobium), differences in the fine mechanisms regulating N2 fixation exist and have been reviewed (Fischer 1994; Kaminski et al. 1998). 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).

Lessons from Genomics Genome size and organization among nitrogen-fixers can vary widely. The genome of the cyanobacteria Nostoc (which is a symbiont of cycads, Gunnera and others) is among the largest from prokaryotes, with nearly 10 Mb (Meeks et al. 2001). In contrast the genome of the Azolla cyanobacterium symbiont has suffered a large reduction (Ran et al. 2010), the true Azolla symbionts are non-culturable. The sequences of the genomes of the legume-nodulating bacteria belonging to the genera of Mesorhizobium, Ensifer and Bradyrhizobium revealed contrasting chromosome sizes and highly diverging genomes. Large genomic differences have been found as well among Frankia genomes from different plant hosts (Normand et al. 2007). 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 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 2000) but has been detected in other arqueae. The genome sequence of the Bacteroidete Candidatus Azobacteroides pseudotrichonymphae from the guts of old termites revealed nitrogen-fixing genes (Hongoh et al. 2008). Genes for nitrogen fixation were discovered in the Fusobacterium nucleatum genome, no members of the Fusobacteria phylum were previously known to fix nitrogen. Similarly the genomes of Candidatus Nitrospira defluvii, of Fibrobacter succinogenes and Deferribacter desulfuricans revealed nitrogen-fixation pathways, for these to our knowledge, there was no evidence yet of nitrogen-fixation activity. These bacteria are the first representatives of their own phylum to be described as nitrogen-fixing species.

Dinitrogen-Fixing Prokaryotes

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The genome analysis of the chlorinated-ethene-respiring Dehalococcoides ethenogenes strain 195 showed the presence of a complete nitrogenase operon and subsequently the nitrogen fixation was demonstrated by incorporation of 15N and transcription of nifD gene (Lee et al. 2009). Interestingly this is a strain that belongs to the phylum Chloroflexi previously known to fix N2. Out of 4 sequenced strains only Dehalococcoides ethenogenes contained nif genes and these resembled Clostridium and archaea nif genes, all anaerobic. Hydrogenobacter thermophilus strain TK-6 belongs to the family Aquificaceae (Yoshino et al. 2001), in a the phylum Aquificae (one of the earliest branching in Bacteria) and has the nifH gene but nifDK are pseudogenes. Non-cultured insect endosymbionts can be studied by genomic aproaches; the reduced genome from the spheroid body from Rhopalodia gibba has revealed its close relatedness to free-living nitrogen-fixing cyanobacterium (Kneip et al. 2008).

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 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.

Conclusions and Perspectives of Application of Nitrogen-Fixation Research

References

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. 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 an ambitious goal is feasible is difficult to predict. The modification of cereals such as rice to render them capable of forming nodules is being explored based on the large knowledge of symbiotic genes in model legume plants (Rolfe et al. 1998). The identification and selection of plant-associated microorganisms and their genetic improvement is an alternative strategy for obtaining agricultural crops that benefit from prokaryotic N2 fixation. N2 fixation from associated bacteria is being considered as a suitable mode to exploit N2 fixation in nonlegumes. 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 and proteomics 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

Acknowledgments Thanks to Julio Martı´nez Romero for technical support. To PAPIIT IN200709 and IN205412 from UNAM.

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12 Magnetotactic Bacteria Dennis A. Bazylinski1 . Christopher T. Lefe`vre2 . Dirk Schu¨ler3 1 School of Life Sciences, University of Nevada at Las Vegas, Las Vegas, NV, USA 2 CEA Cadarache/CNRS/Universite´ Aix-Marseille II, UMR7265 Service de Biologie Ve´ge´tale et de Microbiologie Environnementale, Laboratoire de Bioe´nerge´tique Cellulaire, Saint Paul lez Durance, France 3 Department Biologie I, Ludwig-Maximilians-Universita¨t Mu¨nchen, Planegg-Martinsried, Germany

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Ecology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Detection and Collection of Magnetotactic Bacteria from Natural Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Diversity and Physiology of the Magnetotactic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 456 Alphaproteobacteria Class . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 456 Gammaproteobacteria Class . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Deltaproteobacteria Class . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 458 Nitrospirae Phylum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460 Other Groups . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462 Evolution of Magnetotaxis and the First Magnetosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462 Cultivation and Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 463 Magnetotaxis, Chemotaxis, Aerotaxis, and Phototaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 466 Axial Magneto-aerotaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 468 Polar Magneto-aerotaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 468 Revised Model of Magnetotaxis: Redoxtaxis . . . . . . . . . . . . 469 Phototaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 470 Magnetosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 470 Magnetic and Mineral Properties of Magnetosomes . . . . 471 Arrangement of Magnetosomes Within Cells of Magnetotactic Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 474 Biomineralization of Magnetosomes . . . . . . . . . . . . . . . . . . . . . . 475 The Magnetosome Membrane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 477 Magnetosome Membrane Proteins . . . . . . . . . . . . . . . . . . . . . . 477 Genomics and Genetics of Magnetotactic Bacteria . . . . . . . . 480 Molecular Organization of Magnetosome Genes . . . . . . . 480 Genetic Manipulation of Magnetotactic Bacteria . . . . . . . 483 Applications of Magnetotactic Bacteria, Magnetosomes, and Magnetosome Crystals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 484 Mass Culture of Magnetotactic Bacteria . . . . . . . . . . . . . . . . 484

Applications of Cells of Magnetotactic Bacteria . . . . . . . . 484 Applications of Magnetosomes . . . . . . . . . . . . . . . . . . . . . . . . . . 484 Geological Significance of Magnetotactic Bacteria and Magnetosomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 485 Abstract Prokaryotes that exhibit magnetotaxis, collectively known as the magnetotactic bacteria, are those whose direction of motility is influenced by the Earth’s geomagnetic and externally applied magnetic fields. These ubiquitous, aquatic microorganisms represent a morphologically, phylogenetically, and physiologically diverse group that biomineralize unique organelles called magnetosomes that are responsible for the cells’ magnetotactic behavior. Magnetosomes consist of magnetic mineral crystals, either magnetite (Fe3O4) or greigite (Fe3S4), each enveloped by a phospholipid bilayer membrane that contains proteins not present in other membranes. While there are several different magnetite and greigite crystal morphologies, mature crystals of both minerals are always in the single magnetic domain size range, about 35–120 nm, thus having the highest possible magnetic moment per unit volume. In most magnetotactic bacteria, magnetosomes are arranged as a chain within the cell thereby maximizing the magnetic dipole moment of the cell causing the cell to passively align along magnetic field lines as it swims. Magnetotaxis is thought to function in conjunction with chemotaxis in aiding magnetotactic bacteria in locating and maintaining an optimal position in vertical chemical concentration gradients common in stationary aquatic habitats, by reducing a three-dimensional search problem to one of a single dimension. Although the detection of magnetotactic bacteria in samples collected from natural environments is relatively easy, the magnetotactic bacteria are a fastidious group of prokaryotes and special culture conditions are necessary for their isolation and cultivation. Phylogenetically, most known cultured and uncultured magnetotactic bacteria are associated with the Alpha-, Gamma-, and Deltaproteobacteria classes of the Proteobacteria phylum and the Nitrospirae phylum. All cultured species are either microaerophiles or anaerobes or both. Most cultured species of the Alpha- and Gammaproteobacteria classes are microaerophiles that grow chemolithoautotrophically using reduced sulfur compounds as electron sources and the

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Calvin-Benson-Bassham cycle or the reverse tricarboxylic acid cycle for autotrophy and chemoorganoheterotrophically using organic acids as electron and carbon sources. Those in the Deltaproteobacteria are sulfate-reducing anaerobes that only grow chemoorganoheterotrophically. Almost all cultured species exhibit nitrogenase activity and thus fix atmospheric nitrogen and many denitrify. Magnetotactic bacteria thus show a great potential for iron, nitrogen, sulfur, and carbon cycling in natural environments. Genetic determinants for magnetosome synthesis, the mam and mms genes, are organized as clusters in the genomes of all magnetotactic bacteria examined. These clusters are in close proximity to each other within the genomes and are surrounded by genomic features that suggest that magnetosome genes are organized as a magnetosome gene island that might be transmitted to many different bacteria through horizontal gene transfer. Through the development of genetic systems in some magnetotactic bacteria, the functions of several magnetosome membrane proteins in the biomineralization of the magnetite magnetosome chain have been demonstrated although the roles of most remain unknown. Bacterial magnetosomes have novel physical and magnetic properties and also geological significance and have been used in a large number of commercial and medical applications.

Introduction Magnetotactic bacteria are gram-negative, motile prokaryotes that synthesize intracellular, membrane-bounded crystals of magnetic iron oxide or iron sulfide minerals. The mineral crystals together with their associated membrane are called magnetosomes (Balkwill et al. 1980) and cause the bacteria to orient and swim along external magnetic and geomagnetic field lines. These microorganisms were first described by Salvatore Bellini in 1963 in a publication of the Instituto di Microbiologia of the University of Pavia, Italy (Bellini 1963, 2009a, b). He observed large numbers of bacteria that swam in a consistent, single, northward direction and referred to them as ‘‘batteri magnetosensibili’’ (magnetosensitive bacteria) and speculated that the magnetic behavior of the cells was due to an internal ‘‘magnetic compass.’’ This internal ‘‘magnetic compass’’ was later confirmed by Richard P. Blakemore who independently rediscovered magnetotactic bacteria in 1974 and was the first to observe magnetosomes within cells of these microorganisms (Blakemore 1975). Magnetotactic bacteria are indigenous in sediments or chemically stratified water columns where they occur predominantly at the oxic-anoxic interface/transition zone (OAI or OATZ) and the anoxic regions of the habitat or both. They represent a diverse group of microorganisms with respect to morphology, phylogeny, and physiology. Despite the efforts of a number of different research groups, few representatives of this group of bacteria have been isolated in axenic culture since their discovery, and even fewer have been adequately described in the literature. Therefore, little is known about their metabolic plasticity, whereas their diverse morphology and phylogeny has

been analyzed to some extent by culture-independent methods. To date, the only validly described species of magnetotactic bacteria are members of the genus Magnetospirillum. Representatives of this genus have been isolated reproducibly from various aquatic environments, can now be grown relatively easily in mass culture, and are genetically tractable. Thus, much of the knowledge regarding the metabolism, genetics, and biochemistry of magnetotactic bacteria is derived from studies involving strains of this genus.

Ecology Magnetotactic bacteria are cosmopolitan in distribution and ubiquitous in sediments of freshwater, brackish, marine, and hypersaline habitats as well as in chemically stratified water columns of these environments (Bazylinski and Frankel 2004). They have also been found in some wet soils (Fassbinder et al. 1990) although it is not known whether their presence is common in them. The occurrence of magnetotactic bacteria appears to be dependent on the presence of an oxic-anoxic interface (OAI) that represents opposing gradients of oxygen from the surface and reduced compounds (usually reduced sulfur species) in sediments or water columns. Generally, the highest numbers of magnetotactic bacteria are observed at the OAI of sediments or chemically stratified water columns (Moskowitz et al. 2008). Moreover, within the OAI, different species of magnetotactic bacteria occupy different positions that are also probably dependent on specific chemical conditions. Magnetotactic bacteria are known to biomineralize two magnetic minerals: the iron oxide magnetite (Fe3O4) or the iron sulfide greigite (Fe3S4). Most magnetotactic bacteria produce only one mineral although there is a group that synthesizes both. Typically, the magnetiteproducers are found at the OAI proper while the greigiteproducers are found below the OAI when the anoxic zone is sulfidic (Moskowitz et al. 2008). Magnetotactic bacteria can thus be considered as typical examples of gradient organisms. For many years, magnetotactic bacteria were thought to be restricted to habitats with pH values near neutral and at ambient temperature. Very recently, however, Lefe`vre et al. described an uncultured, moderately thermophilic magnetotactic bacterium in hot springs in northern Nevada (Lefe`vre et al. 2010b) with a probable upper growth limit of about 63  C. In addition, this same group isolated several strains of obligately alkaliphilic magnetotactic bacteria from different aquatic habitats in California including the hypersaline, extremely alkaline Mono Lake (Lefe`vre et al. 2011b). These strains have an optimal growth pH  9.0. None yet have been found in habitats that are strongly acidic (e.g., acid mine drainage).

Detection and Collection of Magnetotactic Bacteria from Natural Environments The detection of magnetotactic bacteria in environmental water and sediment samples is relatively easy due to their magnetic behavior in turn due to their permanent magnetic dipole

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. Fig. 12.1 Image of the ‘‘hanging drop’’ setup used for the detection of magnetotactic bacteria in water and sediment samples (Schu¨ler 2002). A drop of water/sediment is placed on a cover slip and inverted and then set on a small rubber o-ring on a microscope slide. The south pole (S) of a bar magnet is placed next to one side of the drop, here the right side. North-seeking magnetotactic bacteria will swim to the right edge of the drop (shown at arrow) and accumulate making them easy to detect and observe microscopically. Using this technique, perturbation of the drop by air currents and evaporation of the drop is reduced. In addition, sediment in the drop settle to the drop’s lowest point, leaving the edge of the drop clear to view the bacteria. The hanging drop was used in the video > Figs. 12.5, > 12.12, > 12.13, and > 12.15

moment. A simple method is to put a drop of water or sediment on a microscope slide which is then set on the microscope stage. A bar magnet is now placed on the microscope stage near the drop so the axis of the magnet is parallel to the plane of the slide and passes through the center of the drop. The magnetic field at the drop should be at least a few gauss and the bar magnet should be oriented so that the ‘‘south’’ magnetic pole (the pole that attracts the north end of a magnetic compass needle) is on the microscope stage in such a way that all the magnetotactic bacteria are guided in one direction until they reach and accumulate at the edge of the drop of water and/or sediment where they can be observed. If the magnet is rotated 180 , the bacteria will also rotate and swim away from the edge of the drop. The use of phase contrast or differential interference contrast microscopy provides much better contrast than bright field microscopy making cells much easier to observe. A commonly used modification of the procedure described above is the so-called hanging drop technique in which a drop of water/sediment is placed on a cover slip and inverted and then set on a small rubber o-ring on the microscope slide (> Fig. 12.1; Schu¨ler 2002). This technique eliminates perturbation of the drop by air currents and reduces evaporation of the drop. It also allows sediment in the drop to settle to the drop’s lowest point, leaving the edge of the drop clear to view the bacteria. Both procedures work well if there are good concentrations of magnetotactic bacteria in the samples. To ensure visualization of cells if concentrations are low, one can magnetically enrich for higher numbers of cells by placing the south pole (in the Northern Hemisphere; the north pole of the magnet is used in the Southern Hemisphere) of a bar magnet adjacent to the outer wall of a jar filled with sediment and water. If magnetotactic

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. Fig. 12.2 Magnetic enrichment of magnetotactic bacteria from a water and sediment sample in a jar by applying the south pole of a magnet (M) outside the jar several centimeters above the water-sediment interface for about 30 min. Magnetotactic bacteria that accumulate at the magnet are shown as a dark spot at the arrow. Interestingly, although one would expect only north-seeking magnetotactic bacteria to accumulate near the magnet, those of both polarities collect near the magnet

bacteria are abundant in the sample, a brownish or grayish to white spot consisting mainly of magnetotactic bacteria will form next to the inside of the glass wall closest to the south pole of the bar magnet (> Fig. 12.2). Cells can be easily removed from the jar with a Pasteur pipette and examined as described above. An extension and scale-up of the magnetic collection method was recently described (Jogler et al. 2009b). By using larger ‘‘magnetic traps’’ holding up to several liters of sediment slurry, large numbers (about 108 cells per liter of sediment) of diverse uncultivated magnetic cells can be selectively harvested from large volumes of sediment samples. In this method, bacteria are magnetically directed toward the tips of collection tubes, from which they can be conveniently collected. Magnetotactic bacteria commonly enrich (increase in numbers) in sediment samples in jars or aquaria stored in dim light at room temperature for several weeks to months. In several studies, successions of different magnetotactic bacterial morphotypes have been observed during the enrichment process. Surprisingly, magnetotactic bacteria sometimes remain active for several years in aquaria without addition of nutrients. Characterization of the large ovoid Nitrospirae, Candidatus Magnetoovum mohavensis, was only possible due to its enrichment in samples incubated for several months after collection (Lefe`vre et al. 2011a). It is important to note that all methods commonly used for the detection and collection of uncultivated magnetotactic bacteria are inherently selective for cells which are highly motile, abundant, and at least temporarily tolerate exposure to atmospheric concentrations of oxygen. Thus, modifications to these techniques to detect, collect, and cultivate environmental magnetotactic bacteria that are at very low concentrations in the sample, that swim very slowly, or that are poisoned quickly by oxygen potentially may reveal an even greater diversity.

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Diversity and Physiology of the Magnetotactic Bacteria Even before the advent of molecular phylogenetics, the great diversity of magnetotactic bacteria was obvious to most investigators that study them because of the large number of different, sometimes unique, morphotypes observed in environmental samples of water and sediment. Commonly observed morphotypes include coccoid to ovoid cells, rods, vibrios, and spirilla of various dimensions. Two of the more unique morphotypes include a group of apparently multicellular bacteria referred to as MMPs and a very large rod provisionally named Candidatus Magnetobacterium bavaricum. Regardless of their morphology, all magnetotactic bacteria studied thus far are motile by means of flagella and have a cell wall structure characteristic of typical gram-negative bacteria with one exception: uncultured, freshwater magnetotactic bacteria belonging to the Nitrospirae phylum appear to have a more complex cell wall structure (Jogler et al. 2011; Lefe`vre et al. 2011a). 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 within the cell. In the majority of magnetotactic bacteria, the magnetosomes are aligned in chains of various lengths and numbers along the 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 (Moench and Konetzka 1978; Moench 1988; Cox et al. 2002). Besides magnetosomes, large inclusion bodies containing elemental sulfur, polyphosphate, or poly-b-hydroxybutyrate (PHB) are common in magnetotactic bacteria collected from natural environments and in pure culture (Bazylinski et al. 2004; Schultheiss et al. 2005). In cultivated magnetospirilla, PHB granules were found to be associated with phasin-like proteins as in other PHB-producing bacteria (Schultheiss et al. 2005). The most commonly observed type of magnetotactic bacteria present in environmental samples are coccoid to ovoid cells, the so-called magnetococci, possessing two flagellar bundles on one somewhat flattened side. This bilophotrichous type of flagellation resulted in the provisional name ‘‘Bilophococcus’’ for the genus of these bacteria (Moench 1988). Interestingly, marine magnetococci possess a sheath surrounding their flagellar bundles (Lefe`vre et al. 2010c). Two representative strains of this morphotype are now in axenic culture (Frankel et al. 1997; Lefe`vre et al. 2009); one now named Magnetococcus marinus (Schu¨bbe et al. 2009; Bazylinski et al. 2012a). The phylogenetic diversity of magnetotactic bacteria, including both those in axenic culture and those collected from natural environments, is also considerable and based on the sequence of their 16S rRNA genes. To date, representatives of the magnetotactic prokaryotes are phylogenetically associated with four major lineages within the domain bacteria and three within the Proteobacteria. No magnetotactic bacterium

phylogenetically associated with the Archaea has yet been discovered. Although most known cultured and uncultured magnetotactic bacteria belong to the Alpha-, Gamma-, and Deltaproteobacteria classes of the Proteobacteria phylum, several uncultured species are affiliated with the Nitrospirae phylum and one, assigned strain SKK-01, to the candidate division OP3, part of the Planctomycetes-Verrucomicrobia-Chlamydiae (PVC) bacterial superphylum (Kolinko et al. 2012) (> Fig. 12.3). The physiology of known magnetotactic bacteria, including that determined experimentally with cultured strains and that inferred from uncultured types, is also quite diverse. In general, however, the physiology of magnetotactic bacteria in almost all cases suggests that they are important in the cycling of key elements including iron, sulfur, nitrogen, and carbon in natural habitats.

Alphaproteobacteria Class Magnetotactic bacteria in the Alphaproteobacteria are only known to biomineralize magnetite and include: all cultured species of the freshwater genus Magnetospirillum (Burgess et al. 1993; Schu¨ler et al. 1999); all of the bilophotrichous magnetotactic cocci including the cultured Magnetococcus marinus strain MC-1 (DeLong et al. 1993; Schu¨bbe et al. 2009; Bazylinski et al. 2012a) and strain MO-1 (Lefe`vre et al. 2009) and numerous uncultured types (Spring et al. 1994, 1998; Lin and Pan 2009), the marine vibrios Magnetovibrio blakemorei strains MV-1 and MV-2 (DeLong et al. 1993; Bazylinski et al. 2012b), and the marine spirilla Magnetospira thiophila strain MMS-1 and strain QH-2 (Williams et al. 2012; Zhu et al. 2010). Using in situ hybridization with fluorescently labeled oligonucleotide probes, it was demonstrated that members of the Alphaproteobacteria class represent the dominant proportion of uncultured magnetotactic bacteria in many freshwater and marine environments (Spring et al. 1992, 1994, 1998). Because many uncultured magnetotactic Alphaproteobacteria contain intracellular sulfur globules (Moench 1988; Cox et al. 2002), autotrophy and/or mixotrophy based on the oxidation of reduced sulfur compounds is thought to be a common feature of these organisms. The ability to fix atmospheric nitrogen was found in all those tested (Bazylinski and Williams 2007). All cultured magnetotactic Alphaproteobacteria are obligate microaerophiles or anaerobes or both (Bazylinski and Frankel 2004; Bazylinski and Williams 2007). Those that tolerate relatively high concentrations of oxygen do not synthesize magnetite under these conditions. They are mesophilic with regard to growth temperature, and none grow much higher than 30  C. Magnetospirillum species have a respiratory form of metabolism and are chemoorganoheterotrophic using organic acids as a source of carbon and electrons (Schleifer et al. 1991). M. gryphiswaldense is also capable of autotrophic and mixotrophic growth using reduced sulfur compounds as a source of electrons (Geelhoed et al. 2010). Although the pathway of autotrophy was not determined, it seems likely that carbon dioxide fixation occurs through the

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. Fig. 12.3 Phylogenetic distribution of cultured and some uncultured magnetotactic bacteria in the Alpha-, Gamma-, and Deltaproteobacteria classes of the Proteobacteria phylum, the Nitrospirae phylum and the candidate division OP3. Magnetotactic bacteria are in bold. The uncultured organisms include: Candidatus Magnetoglobus multicellularis and Ca. Magnetomorum litorale, both forms of the MMP, of the Deltaproteobacteria; all strains in the Nitrospirae; and SKK-01 of the candidate division OP3. All others are cultured

Calvin-Benson-Bassham cycle since a form II ribulose-1,5bisphosphate carboxylase/oxygenase (RubisCO) gene was found in the genome of M. magnetotacticum (Bazylinski et al. 2004). While most species are facultative anaerobes that utilize nitrate as an alternate terminal electron acceptor to oxygen, M. magnetotacticum appears to be an obligate microaerophile that requires oxygen even when growing with nitrate (Bazylinski and Blakemore 1983a; Blakemore et al. 1985). In Magnetospirillum species, magnetite synthesis only occurs at very low levels of oxygen or under anaerobic conditions when nitrate is the alternate terminal electron acceptor to oxygen (Bazylinski and Blakemore 1983a; Blakemore et al. 1985; Schu¨ler and Baeuerlein 1998; Heyen and Schu¨ler 2003). All three described species of Magnetospirillum show dinitrogendependent growth and show nitrogenase activity (the reduction of acetylene to ethylene in nitrogen-free medium) demonstrating that they are capable of nitrogen fixation (Bazylinski and Blakemore 1983b; Bazylinski et al. 2000). In further support

of this, a full series of nif genes is present in the genomes of M. magnetotacticum and M. magneticum. The marine vibrio, Magnetovibrio blakemorei strain MV-1, also has a respiratory metabolism using oxygen, nitrate and nitrous oxide as terminal electron acceptors (Bazylinski et al. 1988, 2012b). It grows chemoorganoheterotrophically with organic and some amino acids as carbon and electron sources (Bazylinski et al. 1988, 2012b; Bazylinski and Williams 2007) and also chemolithoautotrophically using reduced sulfur compounds as an electron source (Bazylinski et al. 2004, 2012b). This strain utilizes the Calvin-Benson-Bassham cycle for autotrophy: cell-free extracts display RubisCO activity and the strain possesses a form II 326 RubisCO gene (Bazylinski et al. 2004). MV-1 also grows chemoorganoautotrophically with formate as the electron donor (Bazylinski et al. 2004, 2012b). This strain shows nitrogenase activity under both heterotrophic and autotrophic conditions (Bazylinski and Williams 2007; Bazylinski et al. 2012b).

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The cultured marine spirilla, Magnetospira thiophila strain MMS-1 and strain QH-2, both appear to be obligate microaerophiles that grow with organic acids as carbon and electron sources (Williams et al. 2012; Zhu et al. 2010). Chemolithoautotrophic growth is also supported in M. thiophila by thiosulfate but not sulfide (Bazylinski and Williams 2007). This species also displays nitrogenase activity under heterotrophic and autotrophic conditions (Bazylinski and Williams 2007). The known cultured and uncultured magnetotactic cocci are not closely related to other Alphaproteobacteria and form their own clade within the Alphaproteobacteria that is basal to the rest of the group (> Fig. 12.3). Many uncultured magnetotactic cocci contain sulfur globules even when sulfide is not apparent or measureable in the sample from which they were collected (Moench 1988; Cox et al. 2002) suggesting an autotrophic or mixotrophic metabolism based on the oxidation of reduced sulfur compounds. The two cultured magnetococci, Candidatus Magnetococcus marinus and strain MO-1, are obligately microaerophilic and grow autotrophically on sulfide and thiosulfate (Williams et al. 2006; Lefe`vre et al. 2009). Ca. Magnetococcus marinus utilizes the reverse (reductive) tricarboxylic acid cycle for carbon dioxide fixation and autotrophy (Williams et al. 2006). It also grows chemoorganoheterotrophically with acetate as the carbon and electron source and is capable of nitrogen fixation based on the strain exhibiting nitrogenase activity and the presence of a full suite of nif genes in its genome (Bazylinski and Williams 2007; Schu¨bbe et al. 2009).

Gammaproteobacteria Class Simmons et al. (2004) provided some evidence for a putative greigite-producing rod belonging to the Gammaproteobacteria. However, the mineral present in its magnetosomes was never identified and a thorough examination of the phylogenetic relationship of this organism raised doubts to its affiliation with this group (Amann et al. 2007). Only recently have magnetotactic bacteria, specifically two strains designated BW-2 and SS-5, been reported to unequivocally belong to the Gammaproteobacteria (Lefe`vre et al. 2012), and thus, there is little information regarding the extent of the diversity of magnetotactic bacteria in this class. Both organisms are mesophilic, microaerophilic rods and biomineralize magnetite. BW-2 and SS-5 are not closely related: BW-2 belongs to the Thiotrichales order whereas SS-5 to the Chromatiales. Very recently, a large group of uncultured and one cultured greigite-producing rods were found to be phylogenetically affiliated with the Deltaproteobacteria (Lefe`vre et al. 2011d). Strain BW-2 was isolated from sediment and water collected from a brackish, sulfidic spring at Badwater Basin at Death Valley, California in which the dominant magnetotactic bacteria were greigite-producing rods (Lefe`vre et al. 2012). Cells are motile by a single polar, unsheathed bundle of seven flagella. This strain is only known to grow chemolithoautotrophically

using sulfide and thiosulfate as electron donors. Cells produce intracellular sulfur globules, and thiosulfate is oxidized completely to sulfate. Cells show nitrogenase activity. Strain SS-5 was isolated from sediment and water collected from the southeastern shore of the hypersaline Salton Sea, California (Lefe`vre et al. 2012). Cells possess a single polar flagellum. Like those of BW-2, cells grow chemolithoautotrophically with sulfide and thiosulfate (oxidized completely to sulfate) but also show potential for heterotrophic growth on succinate. Although they do not produce intracellular sulfur globules, they synthesize large deposits of phosphate-rich inclusions. Unlike all magnetotactic bacteria tested, SS-5 did not show nitrogenase activity.

Deltaproteobacteria Class The Deltaproteobacteria contain both magnetite- and greigiteproducing magnetotactic bacteria and include: the various forms of the uncultured MMP which biomineralize either or both minerals (DeLong et al. 1993; Keim et al. 2003; Abreu et al. 2007; Simmons and Edwards 2007); a group of uncultured and two cultured (strains BW-1 and SS-2), large, rod-shaped bacteria that biomineralize either or both minerals (Lefe`vre et al. 2011d); the magnetite-producing, rod-shaped, sulfate-reducer Desulfovibrio magneticus strain RS-1 isolated from a freshwater river in Japan (Sakaguchi et al. 1993, 2002); and several similar strains of obligately alkaliphilic, sulfate-reducing, magnetiteproducing rods isolated from extremely alkaliphilic habitats in California, USA, that, based on 16S rRNA gene sequence identity, likely represent new strains of Desulfonatronum thiodismutans, a known non-magnetotactic Deltaproteobacterium (Lefe`vre et al. 2011b). All magnetotactic Deltaproteobacteria are mesophilic based on their growth temperature or the temperature of their habitats. The MMP. One of the most interesting and unusual examples of prokaryotic morphology is that of the organisms known as magnetotactic multicellular prokaryotes (MMPs; also known as the magnetotactic multicellular aggregate (MMA) (Farina et al. 1983; Lins and Farina 1999), the magnetotactic multicellular organism (MMO) (Keim et al. 2004a), and magnetotactic multicellular bacteria (Shapiro et al. 2011). The acronym MMP originally represented many-celled magnetotactic prokaryote (Rodgers et al. 1990a, b) because it was difficult to prove that the organism was truly multicellular. Because of a number of recent findings suggesting that individual cells interact and/or communicate with each other, many researchers now use MMP for multicellular magnetotactic prokaryote (e.g., Wenter et al. 2009). Three MMPs have been tentatively named: Candidatus Magnetoglobus multicellularis (Abreu et al. 2007), Ca. Magnetomorum litorale (Wenter et al. 2009), and Ca. Magnetananas tsingtaoensis (Zhou et al. 2012). Interestingly, despite their unique morphology, if not for its magnetotactic behavior, it is unlikely that the MMP would have been discovered.

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. Fig. 12.4 Brightfield transmission electron microscopy (TEM) images of the multicellular magnetotactic prokaryote (MMP). (a) Thin-section of an MMP showing its many-celled nature and the acellular compartment in the center of the rosette of cells. Electron-lucent vacuoles may represent poly-b-hydroxybutyrate (PHB) granules (Micrograph courtesy of F. Abreu) (b) Unstained cells of an MMP revealing the numerous greigite-containing magnetosomes within the organism mostly arranged in short chains. (c) Outer surface of an unstained MMP. Flagella are distributed asymmetrically on each cell and cover the cell on one side

MMPs are relatively large for prokaryotic microorganisms and range from about 3- to 12-mm in diameter (Bazylinski et al. 1990; Rodgers et al. 1990a, b) (> Fig. 12.4). It is best described as an aggregation of about 10–60 g-negative, genetically similar cells that swim only as an intact unit and not as individual cells (Bazylinski et al. 1990; Rodgers et al. 1990a, b; Simmons and Edwards 2007). Cells that become separated from the intact unit die quickly (Abreu et al. 2006). Cells are asymmetrically flagellated, the surface of the cell exposed to the surrounding environment covered with numerous flagella (Rodgers et al. 1990a, b; Silva et al. 2007). Most described MMPs are spherical (Bazylinski et al. 1990; Rodgers et al. 1990a, b; Keim et al. 2004a, 2007; Abreu et al. 2007; Wenter et al. 2009), although some are ovoid or pineapple-shaped in morphology (Lefe`vre et al. 2007; Zhou et al. 2012), and all appear to possess a central, acellular compartment (Keim et al. 2004a, 2007). The MMP divides as aggregates without an individual cell stage (Keim et al. 2004b, 2007; Zhou et al. 2012). MMPs are cosmopolitan in distribution in numerous saline aquatic environments, ranging from brackish to hypersaline (Keim et al. 2004a, b; Abreu et al. 2007; Martins et al. 2009). In all cases, the salinity is due to the input of seawater, and many have considered these organisms indigenous only to marine environments (Simmons and Edwards 2007). Recently, non-magnetotactic forms of MMP (referred to as nMMPs) were found in springs and lakes with relatively low salinities (5–11 ppt) and no marine input (Lefe`vre et al. 2010a). The nMMPs have typical MMP morphology but contain up to 60 cells per aggregate. They are phylogenetically closely related to MMPs (Lefe`vre et al. 2010a). Little is known regarding the physiology but it seems very likely that the MMPs are sulfate-reducing bacteria based on the fact that their closest phylogenetic relatives are sulfate-reducers (DeLong et al. 1993; Simmons and Edwards 2007) and that the genes for

dissimilatory sulfite reductase (dsrAB) and dissimilatory adenosine-50 -phosphate reductase (aprA) were detected in purified MMP samples (Wenter et al. 2009). The magnetic mineral greigite in magnetotactic bacteria was first discovered in MMPs (Farina et al. 1990; Mann et al. 1990b). Since then, they have also been found to contain nonmagnetic precursors to greigite (Po´sfai et al. 1998a, b), magnetite (Zhou et al. 2011, 2012), or both magnetite and greigite magnetosomes (Keim et al. 2003; Lins et al. 2007). The greigite crystals in magnetosomes of MMPs are generally pleomorphic although cuboctahedral, elongated-prismatic, and bullet-shaped particles have been observed (Mann et al. 1990b; Po´sfai et al. 1998a, b; Wenter et al. 2009) (see later section on > ‘‘Magnetosomes’’). Only bullet-shaped magnetite crystals have yet been found in magnetosomes of MMPs (Keim et al. 2003; Lins et al. 2007; Zhou et al. 2011, 2012). Magnetosomes are usually loosely arranged in short chains or clusters in individual cells (Mann et al. 1990b; Po´sfai et al. 1998a, b; Lins et al. 2007; Wenter et al. 2009) although there is a general enough consensus in magnetosome arrangement that there is a magnetic dipole to the entire unit (Bazylinski and Frankel 2000; Wenter et al. 2009). It has been also shown that magnetosome chain polarities of individual cells contribute coherently to the total magnetic moment of the MMP in a highly coordinate fashion, suggesting a remarkable degree of magnetic optimization, which is likely inherited by individual cells by a sophisticated reproduction mechanism (Winklhofer et al. 2007). The total magnetic moments of MMPs from different collecting sites ranged from 5  10 16 to 1  10 15 A m2 for one group (Rodgers et al. 1990a, b) and 9–20  10 15 A m2 for Candidatus Magnetoglobus multicellularis (Perantoni et al. 2009), which are sufficient for an effective magnetotactic response. However, magnetic measurements of greigitecontaining MMPs showed that hysteresis loops of these

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Magnetotactic Bacteria

. Fig. 12.5 Sequence showing the typical ‘‘ping-pong’’ motility of the MMP. For the video, see the online version of The Prokaryotes

organisms are not square indicating that MMPs, unlike magnetotactic bacteria that contain a single chain of magnetite magnetosomes, can be demagnetized (Penninga et al. 1995). The type of magnetotaxis displayed by the MMP appears to be polar (see > ‘‘Magnetotaxis, Chemotaxis, Aerotaxis, and Phototaxis’’ section below), although they have been observed to reverse direction in a magnetic field (it is unknown whether they physically turn around). Under oxic conditions in a uniform magnetic field, the swimming speed of the MMP in the preferred direction averages about 105 mm/s. After reaching the edge of a water drop, they sometimes spontaneously swim in the opposite direction and show short excursions of 100– 500 mm at twice the speed of the forward motion in the opposite direction of their polarity after which they return to the same edge of the drop at a slower speed (Rodgers et al. 1990a, b) as shown in > Fig. 12.5. MMPs exhibit this so-called ‘‘ping-pong’’ motility (Rodgers et al. 1990a, b) in magnetic fields at least several times stronger than the Earth’s magnetic field (~0.5 G) (Greenberg et al. 2005). A detailed study of this behavior in hanging drops (Greenberg et al. 2005) revealed that the outward and return excursions show a uniform deceleration and acceleration, respectively. In addition, the probability per unit time of an MMP undergoing a ping-pong increases monotonically with an increase in the strength of the magnetic field. Outward excursions show an unusual minimum distance distribution, also dependent on the magnetic field strength, in which the higher the field strength, the lower the minimum excursion distance. Desulfovibrio magneticus strain RS-1 is an obligate anaerobe that grows and respires with sulfate and fumarate as electron donors (Sakaguchi et al. 1993, 2002). Like all Desulfovibrio species, cells are curved rods (vibrios) that possess a single polar flagellum and show no potential for autotrophic growth. Small organic molecules and some organic acids support heterotrophic growth in this organism. It is the only known cultured

magnetotactic bacterium to be capable of fermentation: pyruvate is fermented to acetate and hydrogen (Sakaguchi et al. 2002). While magnetotactic bacterium have never been associated with extremophilic conditions, recently, three strains of obligately alkaliphilic, anaerobic, sulfate-reducing, magnetotactic bacteria belonging to the Deltaproteobacteria with optimal growth pH’s of 9.0–9.5 were isolated and grown in axenic culture (Lefe`vre et al. 2011b). All strains biomineralize bullet-shaped crystals of magnetite, are closely related to each other, and appear to be strains of Desulfonatronum thiodismutans, a known alkaliphilic sulfate-reducing bacterium that does not biomineralize magnetosomes (Pikuta et al. 2003) based on their very high sequence identities of their 16S rRNA genes (Lefe`vre et al. 2011b). Like D. thiodismutans, cells are vibrioid to helicoid in morphology and possess a single polar flagellum. All strains grow autotrophically and possibly mixotrophically with hydrogen as electron donor. Formate is also utilized as electron donor. Strain BW-1, recently isolated from a saline spring at Badwater Basin, Death Valley National Park (California), and strain SS-2 isolated from the Salton Sea (California) are two members of a group of large, rod-shaped bacteria that biomineralize greigite and/or magnetite (> Fig. 12.6). BW-1 grows chemoheterotrophically using sulfate as a terminal electron acceptor and produces both minerals, the dominant mineral present being dependent upon culture conditions (e.g., sulfide concentration). The greigite crystals appear to be pleomorphic, while the magnetite crystals are bullet-shaped like those of all other deltaproteobacterial magnetotactic bacteria. Both organisms belong to a previous unrecognized group of sulfate-reducing bacteria in the family Desulfobacteraceae (Lefe`vre et al. 2011d).

Nitrospirae Phylum Thus far, no magnetotactic Nitrospirae have been isolated in axenic culture. However, four different uncultured magnetotactic bacteria phylogenetically associated with this phylum have been described in some detail. The large rod, Candidatus Magnetobacterium bavaricum, is the most studied and was first discovered in sediment samples from Lake Chiemsee and Lake Ammersee in southern Germany (Vali et al. 1987; Petersen et al. 1989). Another magnetotactic Nitrospirae, a small rod-shaped bacterium collected from sediment of the Waller See, Germany, was described by Flies et al. (2005b) and designated strain MHB-1. Recently, Lefe`vre et al. (2010b, 2011a) described two new Nitrospirae: a moderately thermophilic species tentatively named Candidatus Thermomagnetovibrio paiutensis strain HSMV-1 found in brackish hot springs within the Great Boiling Springs geothermal field in Gerlach, Nevada, USA, and a large ovoid-shaped organism tentatively named Candidatus Magnetoovum mohavensis strain LO-1 from freshwater sediments of Lake Mead, Nevada. An organism closely related to strain LO-1, designated MWB-1, isolated from Lake Beihai in Beijing, China, was recently described (Lin et al. 2012). All known

Magnetotactic Bacteria

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. Fig. 12.6 TEM images of the cultured strain BW-1, a deltaproteobacterial magnetotactic bacterium that biomineralizes greigite and magnetite. (a) Brightfield TEM image of a cell of strain BW-1 showing chain of magnetosomes (M) and single polar flagellum (F). (b) Dark field TEM image of magnetosomes in a cell of BW-1. Crystals at arrows are magnetite, the others are greigite

magnetotactic Nitrospirae biomineralize bullet-shaped crystals of magnetite. Candidatus Magnetobacterium bavaricum. This cell morphotype was first observed in samples of littoral sediments collected from Lake Chiemsee and Lake Amersee in southern Germany (Vali et al. 1987; Petersen et al. 1989). Ca. M. bavaricum-like cells have also been found in Brazil (Lins et al. 2000), France (Isambert et al. 2007), and China (Lin et al. 2009; Lin and Pan 2009; Li et al. 2010). Because of its large size, volume (average volume ca. 25.8  4.1 mm3), and relative abundance, Ca. M. bavaricum may account for approximately 30 % of the microbial biovolume in the microaerobic zone of sediments and may, therefore, be a dominant fraction of the microbial community in this zone of Lake Chiemsee (Spring et al. 1993). In addition, 16S rRNA sequences very similar to that of Ca. M. bavaricum (>99 % identity) have been retrieved from a number of freshwater and marine habitats and biological reactor columns (Jogler et al. 2010). Cells of Candidatus Magnetobacterium bavaricum are large rods having dimensions of 1–1.5  6–9 mm and are motile by a single polar tuft of flagella (> Fig. 12.7). Cells contain between 600 and 1,000 magnetosomes that contain bullet-shaped crystals of magnetite that range from 110 to 150 nm in length and are arranged as several braid-like bundles (usually 3–5 per cell) of multiple chains (Hanzlik et al. 1996, 2002; Jogler et al. 2010; Li et al. 2010). Many of the crystals display a kink or hook-like feature. The average total magnetic moment per cell was experimentally determined to be approximately 3  10 14 A m2, which is about an order of magnitude higher than that of most other magnetotactic bacteria. Large amounts of bullet-shaped magnetite crystals have been found in some sediments where Ca. M. bavaricum is present suggesting to some that magnetite from this organism accounts for a large proportion (up to 10 %) of

the total magnetization in these sediments (Petersen et al. 1989; Pan et al. 2005). Candidatus Magnetobacterium bavaricum displays polar magnetotaxis, and in a uniform magnetic field, cells swim forward with an average speed of about 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 (Spring et al. 1993). Because Candidatus Magnetobacterium bavaricum is mainly found in the microaerobic zone (OAI) of sediments and contains sulfur-rich globules, it is thought to be a microaerophilic, sulfide-oxidizing bacterium (Spring et al. 1993; Jogler et al. 2010). In addition, a putative large type IV ribulose-1,5bisphosphate carboxylase/oxygenase (RubisCO) subunit gene was found in a 34 kb genomic region of Ca. M. bavaricum, and although these RubisCO-like proteins do not exhibit RubisCO enzymatic activity (Hanson and Tabita 2001), it may be linked to sulfur metabolism in this organism (Jogler et al. 2010). Another magnetotactic Nitrospirae, a small rod-shaped bacterium collected from sediment of the Waller See, Germany, was described by Flies et al. (2005b) and designated strain MHB-1. This organism is a slow moving, rod-shaped bacterium that contains a single bundle of multiple chains of magnetite magnetosomes whose crystals are also bullet-shaped. The uncultured Candidatus Thermomagnetovibrio paiutensis strain HSMV-1 was found in a series of brackish hot springs with temperatures between 32  C and 63  C within the Great Boiling Springs geothermal field in Gerlach, Nevada USA (Lefe`vre et al. 2010b). Cells are small vibrios with a single polar flagellum. The upper limit of growth of this bacterium is probably around 63  C as it was not present in those springs with higher temperatures.

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. Fig. 12.7 Scanning electron microscope (SEM) and TEM images of cells of Candidatus Magnetobacterium bavaricum. (a) SEM image obtained by simultaneous detection of secondary (yellow) and backscattered electrons (blue). Multiple chains of magnetite crystals are visible (blue). (b) SEM image of a cryofractured cell showing two bundles of magnetosome strands. (c) TEM image of an ultrathin section of highpressure frozen and freeze-substituted cells showing strands of magnetosomes aligned parallel to a tubular filamentous structure (at asterisks). MM magnetosome membrane) (Micrographs courtesy of Gerhard Wanner, LMU Mu¨nchen [with kind permission])

Candidatus Magnetoovum mohavensis strain LO-1 was discovered in freshwater sediments of Lake Mead, Nevada USA (Lefe`vre et al. 2011a). This bacterium is relatively large, ovoid in morphology, has a single polar bundle of sheathed flagella, and biomineralizes braid-like bundles (usually three) of multiple chains of bullet-shaped magnetosomes. Although the organism is likely gram-negative, it appears to have an unusual threelayered cell wall. This organisms may be widely distributed as similar organisms have been observed and collected from freshwater and estuarine environments including the Exeter River, New Hampshire USA (Mann et al. 1987a, b); the Pettaquamscutt Estuary, Rhode Island USA (Bazylinski and Frankel 2003); several sites in Germany (Flies et al. 2005a; Amann et al. 2007); and freshwater lagoons (Jacarepia´ Lagoon, Saquarema, Brazil) and brackish waters (Lagoa de Cima, Rio de Janeiro) in southeastern Brazil (Lins et al. 2000). Like those of Candidatus Magnetobacterium bavaricum, cells of Ca. Magnetoovum mohavensis contain sulfur-rich inclusions suggesting a metabolism based on the oxidation of reduced sulfur compounds. The distribution of cells in a natural microcosm was also similar to that found for Ca. Magnetobacterium bavaricum (Jogler et al. 2010) in that the majority of cells were found at the OAI and the upper layer of the anaerobic zone. In semi-solid oxygen-gradient medium, however, cells immediately migrated to the anoxic zone and remained viable for several days. These results indicate that LO-1 is likely an anaerobe that tolerates low concentrations of oxygen. These studies also suggest that this organism is mesophilic. A magnetotactic bacterium morphologically similar to Ca. Magnetoovum mohavensis, strain MWB-1, was isolated from Lake Beihai in Beijing, China, and shares 95% 16S rRNA gene sequence identity with Ca. Magnetoovum mohavensis

(Lin et al. 2012). The watermelon-shaped MWB-1 appears to account for more than 10 % of the natural remanent magnetization of the surface sediment of Lake Beihai (Lin et al. 2012).

Other Groups Recently, by using single cell-based techniques, a magnetotactic bacterium of low abundance was found to belong to the candidate OP3 division of bacteria which so far lacks any cultured representatives (Hugenholtz et al. 1998), based on 16S and 23S rRNA gene sequences (Kolinko et al. 2012). This might indicate that the diversity and phylogenetic distribution of magnetotactic bacteria is underestimated and may extend toward other unrecognized groups.

Evolution of Magnetotaxis and the First Magnetosomes The early initial discovery that greigite- and magnetiteproducing magnetotactic bacteria were affiliated with two evolutionary distinct lines of descent, the Deltaproteobacteria and the Alphaproteobacteria, respectively, led DeLong et al. (1993) to suggest that the trait of magnetotaxis based on iron sulfide and iron oxide had multiple evolutionary origins. At the present, however, considering the now considerable genomic and new phylogenetic information, it seems more likely that the magnetotactic trait is monophyletic originating from a common ancestor regardless of magnetosome mineral composition (Abreu et al. 2011; Jogler et al. 2011) and that it has been passed to many

Magnetotactic Bacteria

diverse prokaryotes through horizontal gene transfer (discussed later in the chapter). Strong evidence for this hypothesis was recently found by the discovery of putative magnetosome (mam) genes in uncultivated MMP from the Deltaproteobacteria (Abreu et al. 2011) and in Candidatus Magnetobacterium bavaricum belonging to the deeply branching Nitrospirae phylum (Jogler et al. 2011), which are homologous to those previously found in the remotely related magnetotactic Alphaproteobacteria. Interestingly, there is a strong correlation with the phylogeny of magnetotactic bacteria and the composition and morphology of the magnetosome mineral crystal they produce (Abreu et al. 2011; Lefe`vre et al. 2011c, 2012). Magnetotactic Alphaproteobacteria only biomineralize morphologically consistent, welldefined crystals of magnetite that include cuboctahedra and elongated prisms (appear as rectangular in projection in electron micrographs). The only two known magnetotactic Gammaproteobacteria also synthesize these types of magnetite crystals. While magnetotactic Deltaproteobacteria biomineralize magnetite or greigite or both, the magnetite crystals are always bulletor tooth-shaped and show much more morphological variation and defects (e.g., kinks) than those produced by the Alphaproteobacteria. The magnetotactic Nitrospirae are only known to biomineralize magnetite crystals whose morphologies are very similar, if not identical, to those found in the Deltaproteobacteria. The great variation in the magnetite crystals of these latter two groups suggests that there is less control over the biomineralization process in these organisms which may be due to the fact that species in these groups appear to possess less magnetosome genes than those in the Alphaproteobacteria (Lefe`vre et al. 2012). Because of this and the fact that the Nitrospirae and the Deltaproteobacteria are the more deeply branching phylogenetic groups, it has been suggested that bullet-shaped magnetite crystals might represent the first magnetosome mineral phase (Abreu et al. 2011).

Cultivation and Isolation Magnetotactic bacteria have generally proven to be fastidious with respect to growth, and the inability to isolate new strains of magnetotactic bacteria and the lack of specific enrichment and isolation media for them have frustrated potential and current researchers in this area for many years. This is in part because these organisms are ubiquitous in freshwater and marine habitats and because many different cell morphotypes can be present in relatively high numbers in collected samples that can be easily visualized. Moreover, many morphotypes actually enrich in samples of mud and water collected in jars or in microcosms that are simply exposed to dim light and left at room temperature without special treatments such as the addition of nutrients (Blakemore 1982; Flies et al. 2005a). Lastly, based on their ecology and those species already in culture, magnetotactic bacteria are clearly gradient-loving or gradient-requiring organisms (e.g., Frankel et al. 1997). Phylogeny of specific morphotypes of magnetotactic bacteria might provide clues as

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to their physiology which might be helpful in their isolation and axenic culture. For example, the phylogeny of the MMPs strongly suggests that these organisms are anaerobic sulfatereducing bacteria (DeLong et al. 1993; Abreu et al. 2007; Simmons and Edwards 2007; Wenter et al. 2009). It is risky, however, to infer physiological capabilities solely on the basis of phylogenetic affiliation. A major problem in the isolation of magnetotactic bacteria is the lack of an effective enrichment medium. Several media have been constructed for the isolation of magnetotactic bacteria, and most that have proven successful are based on the formulation of Blakemore et al. (1979) for freshwater magnetotactic spirilla. While cells of most magnetospirilla grow well in this medium, the medium does not enrich for them when water and/or mud samples containing magnetotactic bacteria are used as an inoculum as non-magnetotactic organisms rapidly outcompete them. However, magnetotactic bacteria can be quite efficiently and selectively separated from sediment particles and contaminating microorganisms by making use of their active magnetically directed motility. Schu¨ler et al. (1999) developed an improved technique for the enrichment and isolation of magnetotactic spirilla by exploiting their magnetotactic behavior, the idea using magnetotactic bacterial cells as inocula that were rendered free of non-magnetotactic contaminants by magnetically separating them using the magnetic capillary ‘‘racetrack’’ of Wolfe et al. (1987). In this technique (modified slightly from the original), a Pasteur pipette is sealed at its thin end in a flame and a cotton plug set at where the wide-mouthed end of the pipette tapers to the thin portion (> Fig. 12.8). The pipette is sterilized, after which the sealed end is filled with filtersterilized (0.2 mm) water from the original sample until the cotton plug is wetted. Sediment and/or water containing magnetotactic bacteria are placed on top of the sterile, wetted cotton plug in the wide-mouthed end of the pipette. The south pole of a bar magnet is placed near the sealed tip of the capillary furthest from the reservoir, and the north pole of an additional bar magnet set near the entrance of the wide-mouthed end of the pipette. Migration to and accumulation of magnetotactic cells at the end of the capillary can be observed by using a dissecting microscope with the lighting set up for dark field. Generally, most fast-swimming cells of magnetotactic bacteria (cocci) will reach the sealed tip in about 20–30 min. When enough cells have accumulated for a reasonable inoculum, the tip of the pipette is broken off and the cells are removed aseptically using a thin syringe needle. The cells are then subsequently transferred to appropriate enrichment media. Although the magnetic capillary racetrack method is quite useful for the separation of larger, faster swimming magnetotactic bacteria such as some large spirilla and the ubiquitous magnetotactic cocci, it can take much longer periods of time for smaller, slower swimming organisms (e.g., cells Magnetovibrio blakemorei strain MV-1) to reach the sealed end of the pipette. Moreover, it is difficult to determine whether these small cells are present at the end of the capillary using a dissecting microscope. After about 30 min, it is not uncommon for motile non-magnetotactic contaminants to reach the end of the sealed capillary. Although this technique has

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. Fig. 12.8 Image of the magnetic ‘‘racetrack’’ described by Wolfe et al. (1987). In this technique (modified slightly from the original), a Pasteur pipette is sealed at its thin end in a flame and a cotton plug placed where the wide-mouthed end of the pipette tapers to the thin portion. The pipette is sterilized (by autoclaving) after which the sealed end is filled with filter-sterilized (0.2 mm) water from the original sample until the cotton plug is wetted. Sediment and/or water containing magnetotactic bacteria are placed on top of the sterile, wetted cotton plug in the wide-mouthed end of the pipette. North-seeking cells are purified by placing the south pole of a bar magnet near the sealed tip of the capillary furthest from the reservoir and the north pole of an additional bar magnet set near the entrance of the wide-mouthed end of the pipette thereby directing north-seeking cells to the sealed end of the capillary containing sterile water. Accumulation of magnetotactic cells at the end of the capillary can be observed by using a dissecting microscope with the lighting set up for dark field. Once enough cells have accumulated at the tip, the sealed pipette tip is broken off and cells are removed aseptically using a thin syringe needle

proven effective in a number of studies, it does not guarantee a homogenous population unless only one type of magnetotactic bacterium is present in the original sample. There is a report questioning whether cells purified by this technique reflect the diversity of magnetotactic bacteria in the original environmental samples (Lin et al. 2008). This should be assumed considering the very diverse swimming speeds of different magnetotactic bacteria. This limitation of magnetic collection can be circumvented by the application of single cell techniques, such as microscopically controlled micromanipulation and cell sorting, by which any conspicuous morphotype can be targeted and separated from mixed environmental communities of magnetotactic bacteria (Kolinko et al. 2012). All known magnetite-producing magnetotactic bacteria are microaerophiles (atmospheric oxygen concentrations are inhibitory to growth), anaerobes, or facultatively anaerobic microaerophiles. Most media used for the growth of these organisms are semi-solid oxygen concentration gradients or liquid anaerobic media. In general, relatively low concentrations of nutrients appear more favorable for the isolation of magnetotactic bacteria compared to richer media containing higher concentrations of carbon and nitrogen sources. Although some species, including Desulfovibrio magneticus and some greigite-producing species (e.g., strain BW-1), are obligate anaerobes, most magnetotactic bacteria tolerate short exposures to oxygen during magnetic purification and inoculation, making the strict exclusion of oxygen during cell manipulations unnecessary. However, it is not clear if this is true for all other uncultivated species, and the strict exclusion of atmospheric oxygen from all sampling, enrichment, and cultivation steps wherever possible might increase the success of isolation. Magnetite-producing magnetotactic bacteria are not only sensitive to high concentrations of oxygen but are also redoxsensitive, that is, they do not grow from small inocula in growth medium without the addition of a reducing agent. Thus,

formation of the oxygen/redox gradient in the growth medium is crucial for the growth of magnetite-producing magnetotactic bacteria. Redox buffering by the addition of reducing agents such as sodium thioglycolate, sodium sulfide, ascorbic acid, or cysteine at concentrations of 0.1–0.4 g L 1 or dithiothreitol at 1 mM to the medium is required for growth of these microaerophilic or anaerobic species (Bazylinski et al. 1988; Schu¨ler et al. 1999). The inclusion of resazurin, a redox indicator that is colorless when fully reduced, is very helpful in the determination of whether a liquid growth medium is totally reduced or whether an oxygen concentration-redox gradient has formed in semi-solid medium. In the latter case, the surface of the medium should be oxidized and pink and the anoxic zone at the lower part of the tube should be reduced and colorless. The formation of semi-solid media containing inverse oxygen and sulfide concentration double gradients has been used for the successful enrichment of freshwater and marine magnetotactic bacteria (Bazylinski and Williams 2007; Schu¨ler et al. 1999). The formulation for this gradient medium, a modification of the medium originally developed by Nelson and Jannasch (1983) for the enrichment and isolation of the microaerophilic, filamentous, sulfide-oxidizer Beggiatoa, is described in detail in Schu¨ler et al. (1999). In this medium, the sulfide gradient is the result of the diffusion from sulfide from a solid sulfide agar plug at the bottom of the tube. Growth of magnetite-producing species in all oxygen concentration growth medium initially occurs as a well-defined microaerophilic band of cells at the OAI (the pink: colorless interface) (> Fig. 12.9). As the band thickens and number of cells in the band increases, cells deplete oxygen at the OAI and the band of motile cells moves toward the surface. Many magnetite-producing magnetotactic bacteria are heterotrophic but facultatively chemolithoautotrophic (Bazylinski et al. 2004; Williams et al. 2006; Geelhoed et al. 2010) or are obligately chemolithoautotrophic (Lefe`vre et al. 2012). Oxygen concentration gradient medium can be used for both

Magnetotactic Bacteria

. Fig. 12.9 Growth of the magnetite-producing microaerophilic magnetotactic bacterium strain BW-2 in semi-solid oxygen concentration gradient medium. Cells initially grow as a microaerophilic band of cells at the oxic-anoxic transition zone (OATZ; also known as the oxic-anoxic interface (OAI)) which here is shown as the pink:colorless interface (tube on the left). As the band thickens and number of cells in the band increases, cells deplete oxygen at the OAI and the band of motile cells moves toward the surface. The tube on the right is uninoculated and the OATZ will gradually move downward in the tube as the medium oxidizes

chemolithoautotrophic and chemoorganoheterotrophic growth. For the former, bicarbonate must be included in the medium and all organic compounds should not be present with the possible exception of the reducing agent (e.g., cysteine) and vitamins if required. The best known electron donors for this medium are sulfide and thiosulfate. Sulfide can be supplied as a solid agar plug as described earlier or directly as a sterile solution after autoclaving (Lefe`vre et al. 2012). A problem here is that bands of elemental sulfur often form in this medium as oxygen chemically oxidizes the sulfide over time. In growth medium used to confirm chemolithoautotrophic growth, highly purified agar should be used such as Agar Noble (Difco) or even high-quality agarose because many typical agars contain impurities that might be inhibitory to autotrophic growth. For chemoorganoheterotrophic growth, a carbon source is required and the best choices are organic acids (e.g., succinate, malate) and some amino acids as no magnetotactic bacterium has been

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shown to utilize any other type of organic compound as a carbon source. The concentration of the carbon source should initially be kept low (2 mM) to allow magnetotactic bacteria to compete with possible contaminants (Schu¨ler et al. 1999). Only recently has a greigite-producing magnetotactic bacterium been grown in axenic culture. Strain BW-1 was isolated from a saline spring at Badwater Basin in Death Valley National Park (California) (Lefe`vre et al. 2011d). Cells of this organism were magnetically separated using the magnetic racetrack as described earlier and inoculated into different types of growth medium. BW-1 appears to be an obligate, sulfate-reducing, chemoorganoheterotrophic anaerobe. Interestingly, BW-1 biomineralizes both magnetite and greigite and the proportion of the minerals within magnetosomes appears to be dependent on chemical conditions in the growth medium, for example, the concentration of sulfide (Lefe`vre et al. 2011d). Iron is required for magnetosome synthesis and, therefore, it must be present in the growth medium. The type of iron source is not critical, however, as long as it is kept soluble at neutral pH either by the presence of chelating agents (particularly if the iron is supplied as Fe(III)) or reducing agents which reduce Fe(III) to the much more soluble Fe(II). Ferrous or ferric salts at concentrations between 20 and 50 mM are generally sufficient to allow for both growth and magnetosome formation (Schu¨ler and Baeuerlein 1996, 1998)), which match the concentration range of free soluble iron found in environmental sediment horizons where magnetic bacteria are most abundant (Flies et al. 2005a). Remarkably, the growth of cultivated Magnetospirillum species is inhibited at iron concentrations of >200 mM (Schu¨ler and Baeuerlein 1996), indicating that intracellular magnetite biomineralization is not an adaptation specific to iron-rich environments. Ferric citrate and ferric quinate are the most often used iron source for growth and magnetite biomineralization, as they can be easily prepared and autoclaved together with other medium components usually without problems with precipitation (Blakemore et al. 1979; Schu¨ler et al. 1999). It is important to understand that Fe(II) and Fe(III) inverse concentration gradients form in the oxygen concentration gradient medium described in the previous paragraph due to the presence of the chemical reducing agents. The formation of sulfide in anaerobic cultures of sulfate-reducing bacteria appears to pose a problem regarding iron availability to cells for magnetosome formation (Lefe`vre et al. 2011b). Several strains of obligately alkaliphilic, sulfate-reducing magnetotactic bacteria were isolated but displayed a weak or no magnetotactic response apparently due to scavenging of iron by sulfide produced during sulfate reduction resulting in the precipitation of black iron sulfides. To obtain a stronger magnetotactic response, the iron concentration was increased from 20 to 200 mM and the headspace of the cultures purged every other day with oxygen-free argon gas to decrease the concentration of hydrogen sulfide in the cultures (Lefe`vre et al. 2011b). This iron availability issue may be true for other non-alkaliphilic, sulfate-reducing magnetotactic bacteria such as Desulfovibrio magneticus since this organism produces very few magnetosomes when grown anaerobically with sulfate compared to fumarate (Po´sfai et al. 2006).

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For marine strains and those from other saline habitats, the concentration of salts in the growth medium is important. An artificial seawater (ASW) formula that has been used successfully for the isolation of various different morphotypes of marine magnetotactic bacteria can be found in Bazylinski et al. (1994). For magnetotactic bacteria from brackish environments, the salinity can be determined with a hand-held refractometer and the seawater diluted with distilled water to the appropriate salinity. For those from nonmarine saline environments, the same artificial seawater is effective. In the case of salinities higher than that of seawater, using a salt mixture with approximately the same ratio of the salts in the ASW only at higher concentrations seems to work well. Because of the high amounts of magnesium and calcium in these media, the phosphate concentration should be kept low (1 mM) and should be added from a sterile stock solution to the medium after autoclaving to prevent it from precipitating. The media as described above are effective for the growth of magnetotactic bacteria but are not useful for their isolation. Two general methods have been used to isolate magnetotactic bacteria in pure culture. The first involves the formation of individual colonies. This has been achieved using agar plates of appropriate media such as activated charcoal agar (ACA) (Schultheiss and Schu¨ler 2003). This technique has proven effective for the growth of Magnetospirillum species on solid medium. Activated charcoal scavenges and decomposes toxic-free oxygen radicals and peroxides thought to inhibit the growth of many microaerophiles (Hoffman et al. 1983; Krieg and Hoffman 1986). Once inoculated, ACA plates are incubated under microaerobic or anaerobic conditions under special gas mixtures (e.g., 1 % oxygen in nitrogen) or oxygen-free gases depending upon the organism (Schultheiss and Schu¨ler 2003; Dubbels et al. 2004). A second method for obtaining individual colonies is through the use of solid medium in shake tubes (Bazylinski et al. 1988). This is useful for those organisms that will not form colonies on plates. Both oxygen concentration gradient and anaerobic shake tubes (> Fig. 12.10) can be made using air or oxygen-free gas in the headspace, respectively. Using either agar plates or shake tubes, colonies of magnetotactic bacteria are usually brown or black in color due to the formation of magnetite (> Fig. 12.10; Schultheiss and Schu¨ler 2003; Dubbels et al. 2004). For those organisms that do not form colonies on either plates or in shake tubes, pure cultures can be obtained by a repeated series of dilution to extinction in many of the media described here as long as the dominant bacterium present in the original culture is the one targeted for isolation.

Magnetotaxis, Chemotaxis, Aerotaxis, and Phototaxis After Blakemore’s rediscovery of magnetotactic bacteria, he proposed a model to explain the function of the bacterial magnetosome (Blakemore 1975). The model was based on the assumption that all magnetotactic bacteria are microaerophilic and indigenous in sediments. Frankel and Blakemore (1980)

. Fig. 12.10 Oxygen concentration gradient shake tubes of wild-type (tube labeled WT) and a non-magnetotactic mutant (tube labeled P1) of Magnetovibrio blakemorei. The headspace of the tubes is air. Note that colonies form in a band about a centimeter below the meniscus at the oxic-anoxic transition zone (OATZ). Colonies of the wild-type are black due the production of magnetite and mutants that do not biomineralize magnetite (e.g., P1) form cream-to-pink colored colonies

showed that these bacteria passively align and actively swim along the inclined geomagnetic field lines as a result of their magnetic dipole moment. Blakemore referred to this behavior as magnetotaxis and proposed that magnetotaxis helps to guide cells to swim downward to less oxygenated regions of aquatic habitats presumably to the surface of or within sediments. Once cells reached their preferred microhabitat, they would presumably stop swimming and adhere to sediment particles until conditions changed, as for example, when additional oxygen was introduced. This notion was supported by an observed predominant occurrence of north-seeking magnetotactic bacteria (i.e., swim in the direction indicated by the North-seeking pole of a magnetic compass needle) under oxic conditions in the Northern. Hemisphere while in the Southern Hemisphere, southseeking bacteria appear to predominate (Blakemore et al. 1980; Blakemore 1982). Due to the inclination of geomagnetic field lines in both the Northern and Southern Hemispheres and the direction of downward being reversed, magnetotactic bacteria in both hemispheres therefore swim downward toward sediments (Blakemore 1982). Later findings, including the discovery of large populations of magnetotactic bacteria in the water columns of chemically stratified aquatic habitats and the isolation of obligately microaerophilic, coccoid magnetotactic bacterial strains, made it necessary to modify this view of magnetotaxis. The traditional model did not completely explain how magnetotactic bacteria in

Magnetotactic Bacteria

the anoxic zone of a water column benefit from magnetotaxis, nor did it explain how magnetotactic cocci form microaerophilic bands of cells in semi-solid oxygen-gradient medium. Spormann and Wolfe (1984) showed earlier that magnetotaxis is somehow controlled by aerotaxis in some magnetotactic bacteria, but this alone does not help to explain all observed effects of magnetotaxis. More recently, it was demonstrated (using pure cultures of different types of magnetotactic bacteria) that magnetotaxis and aerotaxis work in conjunction in these bacteria (Frankel et al. 1997). The behavior observed in these strains is now referred to as ‘‘magneto-aerotaxis,’’ which appears to be a more accurate description than magnetotaxis. Moreover, ‘‘magnetotaxis’’ is a misleading term (a misnomer) in that cells do not swim toward or away from a magnetic field as the term implies. The traditional model also failed to explain various types of apparently unusual magnetotactic behavior observed by a number of investigators but without recognizing the fundamental differences between these behaviors (Moench and Konetzka 1978; Blakemore et al. 1980; Spormann and Wolfe 1984). Only when distinct morphotypes of magnetotactic bacteria were isolated and grown in pure culture for detailed studies in using thin, flattened capillaries (Frankel et al. 1997), it became clear that two general types of mechanisms were observed, apparently occurring in different bacteria, termed polar and axial. The distinction between polar and axial behavior can be seen by observing cells in wet mounts under oxic conditions using a microscope and a bar magnet of a few gauss parallel to the plane of the slide (> Fig. 12.11). Polar magnetotactic bacteria, particularly the magnetotactic cocci, swim persistently along magnetic field lines without reversing their direction or turning. If the magnetic field is reversed, the bacteria reverse their swimming direction and continue swimming persistently in the same direction relative to the magnetic field. Polar magnetotactic bacteria from Northern Hemisphere habitats appear to predominately swim parallel to the magnetic field, corresponding to northward migration in the geomagnetic field. Bacteria from the Southern hemisphere swim antiparallel to the magnetic field. It was this consistent swimming behavior that led to the discoveries of magnetotactic bacteria by both Bellini and Blakemore (1975). In contrast, axial magnetotactic bacteria, especially the freshwater spirilla grown in liquid culture, orient along magnetic field lines and swim in both directions displaying frequent reversals of swimming direction with some cells accumulating or getting stuck in approximately equal numbers on both the north and south edges of the water drop (> Fig. 12.11a). The distinction between polar and axial magneto-aerotaxis can also be seen in flattened capillary tubes containing suspensions of cells in reduced medium with one or both ends of the capillary tube open. In the first situation, where one end of the capillary is open (the right end of the capillaries > Fig. 12.11b) and the other sealed, a single oxygen concentration gradient forms beginning at the open end of the capillary. Cells of strain MC-1 in these capillaries rotate 180 after a reversal of B, the magnetic field, and the band separates into groups of

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. Fig. 12.11 Two types of magnetotaxis. (a) Depictions of the polar magnetotactic behavior of strain MC-1 and axial magnetotactic behavior of Magnetospirillum magnetotacticum in water drops under oxic conditions on a microscope slide (B, magnetic field; arrow points northward). Cells of strain MC-1 swim persistently parallel to B (north-seeking motility) and accumulate at the edge of the drop. When B is reversed, cells continue to swim parallel to B (north-seeking motility) and accumulate at the other side of the drop. Cells of M. magnetotacticum swim in either direction relative to B and continue to do so when the field is reversed. (b) Illustrations of aerotactic bands of strain MC-1 and M. magnetotacticum in flat glass capillaries. The right ends of the capillaries are open to air and the left ends are sealed. After reversal of B, cells of strain MC-1 rotate 180 and the band separates into groups of cells swimming in opposite directions along B, away from the position of the band before the reversal. A second reversal results in the reformation of a single band. Cells of M. magnetotacticum also rotate 180 but the band of cells remains intact (Figure adapted from Frankel et al. (1997))

cells swimming in opposite directions along B, away from the position of the band before the reversal. A second reversal results in the reformation of a single band. Cells of Magnetospirillum magnetotacticum also rotate 180 in these capillaries, but the band of cells does not separate and remains intact (> Fig. 12.11). In the second situation (not shown), where both ends of the capillary tubes are open, diffusion of oxygen into the ends of the tubes creates an oxygen concentration gradient at each end of the tube, oriented in opposite directions. Polar magnetotactic bacteria incubated in a magnetic field oriented along the long axis of the tube form an aerotactic band at only one end of the tube, whereas axial magnetotactic bacteria form bands at both ends of the tube. Thus, for polar magnetotactic bacteria, the magnetic field provides an axis and

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direction for motility, whereas for axial magnetotactic bacteria, the magnetic field only provides an axis of motility, pointing to different magneto-aerotactic mechanisms occurring in two types of bacteria. Nonetheless, regardless of the form of magneto-aerotaxis, it appears to function in magnetotactic bacteria by aiding them to more efficiently locate and maintain position in an optimal position in chemical concentration (e.g., oxygen) gradients by reducing a three-dimensional search problem to that of a single dimension, that is, once cells are aligned along inclined geomagnetic field lines, they only have to swim up or down (Frankel et al. 1997).

Axial Magneto-aerotaxis Almost all cultured magnetotactic spirilla exhibit axial magnetoaerotaxis (> Figs. 12.11 and > 12.12) after repeatedly grown in liquid media. In most environmental samples, however, magnetospirilla appear to display polar magneto-aerotaxis (see next section), and there is only a single report of magnetospirilla exhibiting axial magneto-aerotaxis in an environmental sample (Spormann and Wolfe 1984). Thus, axial magnetotactic bacteria may represent only a very small fraction of the total count of magnetotactic bacteria in natural samples, although these organisms are harder to detect in wet mounts or hanging drops using a microscope. Cells representing this type of magnetotaxis were referred to as two-way swimmers, because in a homogeneous medium, they swim in either direction along the magnetic field, B (> Fig. 12.12). In the presence of an oxygen concentration gradient, cells swim parallel or antiparallel to B with aerotaxis determining the direction of migration. Therefore, an aerotactic band of cells forms at both ends of the tube in capillaries where both ends are open, whereas cells displaying a polar magnetotaxis form only one band at the end of the tube corresponding to their magnetic polarity. The aerotactic, axial magnetotactic spirilla appear to use a temporal sensory mechanism for oxygen detection as do most microaerophilic bacteria studied so far (Frankel et al. 1998). Changes in oxygen concentration measured during swimming determine the sense of flagellar rotation. Cells moving away from the optimal oxygen concentration consequently reverse their swimming direction. In this model, changes in oxygen concentration are measured within short intervals implying that these bacteria must be actively motile in order to quickly measure and respond to concentration gradients in their habitat. The combination of a passive alignment along geomagnetic field lines with an active, temporal, aerotactic response provides the organism with an efficient mechanism to locate the microoxic or suboxic zone in its habitat. Therefore, the term magneto-aerotaxis is also an appropriate descriptive term for this tactic behavior.

Polar Magneto-aerotaxis The large majority of uncultured, naturally occurring magnetotactic bacteria display polar magnetotaxis most notably

. Fig. 12.12 Sequence showing magnetotactic spirilla displaying axial magnetotaxis. For the video, see the online version of The Prokaryotes

. Fig. 12.13 Sequence showing magnetotactic cocci displaying polar magnetotaxis. For the video, see the online version of The Prokaryotes

the magnetococci (> Figs. 12.11 and > 12.13). However, as indicated in the previous section, polar magnetotaxis has also been observed in several freshly isolated strains of Magnetospirillum from environmental samples (Schu¨ler et al. 1999). Although individual cells swam back and forth, they had a preference for one direction over the other, and the entire population migrated with a predominantly N-seeking polarity and accumulated at one edge of a hanging drop in magnetic fields. Magnetic polarity, however, was lost upon repeated subcultivation of the new isolate under laboratory conditions, presumably due to the absence of a selective pressure for polarity. The following mechanism for polar magnetotaxis was proposed based on experimental data obtained with an axenic

Magnetotactic Bacteria

culture of the marine magnetotactic coccus Candidatus Magnetococcus marinus strain MC-1 (Frankel et al. 1997). These cocci were shown to swim in both directions along a static magnetic field, B, apparently without the need of physically turning around, by reversing the sense of flagellar rotation. It seems that a two-state sensory mechanism determines the sense of flagella rotation leading to parallel or antiparallel swimming along the geomagnetic field lines. Under higher than optimal oxygen tensions, the cell is presumably in an ‘‘oxidized state’’ and swims persistently parallel to B (> Fig. 12.13), that is, downward in the Northern Hemisphere. Under reducing conditions or suboptimal oxygen concentrations, the cell switches to a second state, the ‘‘reduced state,’’ which leads to a reversal of the flagellar rotation and to a swimming antiparallel to B (upward in the Northern Hemisphere). This two-state sensing mechanism results in an efficient aerotactic response, provided that the oxygen-gradient is oriented correctly relative to B, so that the cell is guided in the right direction to find either reducing or oxidizing conditions. This is especially important because adaptation, which would lead to a spontaneous reversal of the swimming direction, was never observed in controlled experiments with the cocci. The redox sensor, which controls this two-state response, might be similar to the FNR (fumarate and nitrate reduction) transcription factor found in Escherichia coli and other bacteria. The FNR factor is sensitive to oxygen and activates gene expression in the reduced state thereby promoting the switch between aerobiosis and anaerobiosis in E. coli (de Graef et al. 1999). The sensory mechanism in the examined magnetotactic cocci is not only affected by oxygen. Cells exposed to light of short wavelengths (500 nm) also showed a response similar to a switch to the ‘‘oxidized state’’ (Frankel et al. 1997).

Revised Model of Magnetotaxis: Redoxtaxis In this section, we extend the current model of magnetoaerotaxis to a more complex redoxtaxis model. In this case, the unidirectional movement of magnetotactic bacteria in a drop of water would be only one aspect of a sophisticated redoxcontrolled response. Many magnetotactic bacteria are now known to be chemolithoautotrophic using reduced sulfur compounds as a source of electrons (Bazylinski et al. 2004; Bazylinski and Williams 2007; Williams et al. 2006; Lefe`vre et al. 2012). Oxygen is the terminal electron acceptor, but it should be stressed that atmospheric levels of oxygen are toxic to these obligate microaerophiles (Bazylinski and Frankel 2004). Thus, in natural environments, energy conservation in these organisms is strongly dependent on the uptake of reduced sulfur compounds which, in many habitats, are present only in deeper regions at or below the OAI due to the rapid chemical oxidation of these reduced chemical species by oxygen or other oxidants in the upper layers. To overcome the problem of separated pools of electron donor and acceptor, several strategies have been developed by sulfide-oxidizing bacteria. Thioploca spp., for example, use nitrate as a terminal electron acceptor, which is stored

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. Fig. 12.14 Hypothetical model of the function of polar magnetotaxis in bacteria (in the Northern hemisphere). Cells are guided along the geomagnetic field lines depending on their ‘‘redox state’’ either downward to the sulfide-rich zone or upward to the microoxic zone, thereby enabling a shuttling between different redox layers

intracellularly (most of the internal space of the cell is vacuolar) to oxidize sulfide and have developed vertical sheaths in which bundles of motile filaments are located. Thioploca is thought to use these sheaths to efficiently move in a vertical direction in the sediment, thereby accumulating sulfide in deeper layers and nitrate in upper layers (Huettel et al. 1996). For some magnetotactic bacteria, it might also be necessary for them to perform excursions to anoxic zones of their habitat in order to accumulate reduced sulfur compounds. The model shown in > Fig. 12.14 illustrates how polar magnetotaxis might help to guide bacteria, depending on their internal redox state, either downward to accumulate reduced sulfur species or upward to oxidize stored sulfur with oxygen. Thus, we hypothesize that magnetotactic bacteria displaying polar magnetotaxis alternate between two internal redox states. The ‘‘oxidized state’’ would result from the almost complete consumption of stored sulfur, the electron donor. In this state, cells seek deeper anoxic layers where they could replenish the depleted stock of electron donor using nitrate or other compounds as alternative electron acceptor. Eventually, they would reach a ‘‘reduced state’’ in which they would have accumulated a large amount of sulfur which cannot be efficiently oxidized under anaerobic conditions leading to a surplus of reduction equivalents. Cells must therefore return to the microoxic zone where oxygen is available to them as an electron acceptor. In addition, low concentrations of oxygen have been shown to be necessary for the synthesis of magnetosomes in some magnetotactic bacteria (e.g., Blakemore et al. 1985). The advantage of polar magnetotaxis is that an oxygen gradient is not necessary for efficient orientation in the anoxic zone, thereby enabling a rapid return of the cell along large distances to the preferred microoxic conditions. A further benefit would be that cells avoid wasting energy by constantly

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moving along gradients but instead can attach to particles in preferred microniches until they reach an unfavorable internal redox state that triggers a magnetotactic response either parallel or antiparallel to the geomagnetic field lines. In any case, greater than optimal concentrations of oxygen would switch cells immediately to an ‘‘oxidized state’’ provoking the typical downseeking response of magnetotactic bacteria observed in oxic water drops under the microscope. This model is summarized in > Fig. 12.14). Although the model of magneto-aerotaxis for some magnetotactic bacteria appears to make sense, there are still many unanswered questions regarding why bacteria are magnetotactic and biomineralize magnetosomes (Frankel and Bazylinski 2004). For example, Simmons et al. (2006) discovered a population of a new magnetotactic bacterial morphotype in the water column of chemically stratified Salt Pond (Woods Hole, Massachusetts USA) whose cells were greater than 90% south-seeking at specific depths. In addition, even the majority of MMPs were south-seeking at certain depths at Salt Pond. Shapiro et al. (2011) also found a majority of south-seeking MMPs in sites at the Little Sippewissett salt marsh (Woods Hole, Massachusetts, USA). Based on the current model of polar magnetotaxis, these organisms would swim southward/upward toward surface waters containing toxic levels of oxygen and would presumably die. In this way, north-seeking bacteria would be selected for and those of the opposite polarity would be selected against. Other significant questions involve the ability of non-magnetotactic mutants of cultured species to form microaerophilic bands of cells in oxygen-gradient medium similar to the wild type and why some cultured species biomineralize far more magnetite under anaerobic conditions when no gradient is present in the medium (e.g., Magnetovibrio blakemorei). It is important to understand that the magnetotaxis model presented above does not preclude other reasons for the organisms’ ability to biomineralize magnetite and/or greigite. While it seems logical that there is a physiological explanation (e.g., magnetite is known to decompose oxygen radicals such as hydrogen peroxide produced during oxygen respiration (Blakemore 1982)), few hypotheses have been put forward and none have been generally accepted.

Phototaxis Some MMPs and nMMPs show a strong negative phototactic response to white light and wavelengths of light 480 nm (Lefe`vre et al. 2010a; Shapiro et al. 2011) (> Fig. 12.15). Because shorter wavelengths of light, 480 nm (blue to violet), are those that generally penetrate the water column the deepest (Braatsch et al. 2004), this negative phototactic response might function similarly to magnetotaxis in that, if light causes MMPs and nMMPs in nature to swim more or less vertically, then, like magnetotaxis (Frankel et al. 1997), it would at least partially reduce a three-dimensional search problem to a onedimensional search problem for an organism that must locate and maintain an optimal position in vertical chemical and redox gradients common in aquatic habitats. Negative phototaxis in

. Fig. 12.15 Sequence demonstrating the negative phototactic response of the nMMPs. Differential interference microscopy of a hanging drop containing nMMPs and magnetotactic rod-shaped bacteria collected from a pool at ambient temperature at the Great Boiling Springs geothermal field in Gerlach, Nevada, demonstrating the negative phototactic response of the nMMPs. The drop is exposed to a magnetic field and initially the microscope is focused at the edge of the north side of the drop. Note the presence of the magnetotactic rod-shaped bacteria that migrated and accumulated at the edge of the drop. After 2 s, the magnetic field is reversed and the magnetotactic bacteria reverse their swimming direction. At 5 s, the field is reversed again. At about 7 s, the point of focus is moved to the opposite, dark side (the south side), far edge of the drop where the nMMPs have accumulated. Note that they move toward the north side of the drop (toward the left) when exposed to the light. The magnetic field direction is consistent throughout this part of the video. nMMPs continue to move away from the light when followed by the light source of the microscope. For the video, see the online version of The Prokaryotes

this case might increase the efficiency of chemotaxis as does magnetotaxis (Frankel et al. 1997). Alternatively, light might simply drive MMPs and nMMPs downward toward anoxic conditions which are likely favorable to them as it has been inferred from phylogenetic data that they are likely sulfatereducing bacteria (DeLong et al. 1993; Simmons and Edwards 2007; Wenter et al. 2009).

Magnetosomes Magnetosomes have an organic membrane and an inorganic mineral phase. The magnetosome mineral phase consists of tens-of-nanometer-sized crystals of an iron oxide and/or an iron sulfide. The mineral composition of the magnetosome in some magnetotactic bacteria is specific enough for it to be likely under genetic control, in that cells of several cultured

Magnetotactic Bacteria

magnetite-producing species still synthesize an iron oxide and not an iron sulfide, even when hydrogen sulfide is present in the growth medium (Meldrum et al. 1993a, b). However, there are some magnetotactic bacteria (in addition to the MMP), specifically a group of large, slow-swimming, rod-shaped types phylogenetically affiliated with the Deltaproteobacteria (Lefe`vre et al. 2011d), that produce both magnetite and greigite magnetosome crystals aligned within the same chain or chains in the cell (Bazylinski et al. 1993b, 1995). There is some evidence to suggest that environmental conditions, that is, whether the cells are under oxic or anoxic conditions, affect what and how much of each mineral is biomineralized in these organisms (Bazylinski et al. 1995; Lefe`vre et al. 2011d).

Magnetic and Mineral Properties of Magnetosomes The size of the magnetosome mineral crystals also appears to be under control of the organism because almost all magnetotactic bacteria contain crystals that display 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-Ricci and Kirschvink 1992; Frankel and Moskowitz 2003). Smaller particles are superparamagnetic at ambient temperature and do not have stable, remanent magnetization. Larger particles tend to form multiple domains, reducing the remanent magnetization. However, exceptionally large magnetite magnetosomes have been observed in some uncultured cocci from the Southern Hemisphere, having dimensions well above the theoretically determined size limits of single domain magnetite (Farina et al. 1994; Spring et al. 1998; McCartney et al. 2001; Lins et al. 2005). Nonetheless, as evidenced by magnetic holography in the transmission electron microscope (TEM), even these large crystals behave as single magnetic domains when they are present in the cell in a chain configuration where they are magnetized by neighboring crystals (McCartney et al. 2001). 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. 12.16). These morphologies and the consistent narrow size range (Devouard et al. 1998) of intracellular magnetosome particles represent typical characteristics 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 (Bazylinski and Frankel 2003). Iron Oxide-Type Magnetosomes. The iron oxide-type magnetosomes consist solely of magnetite, Fe3O4. The particle morphology of the magnetite crystals in magnetotactic bacteria varies but is generally 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 (> Fig. 12.16) (Mann et al. 1990a; Bazylinski et al. 1994).

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They include: (1) roughly cuboidal (cuboctahedral; Balkwill et al. 1980; Mann et al. 1984a, b), (2) parallelepipedal or elongated-prismatic (rectangular in the horizontal plane of projection; Moench and Konetzka 1978; Towe and Moench 1981; Moench 1988; Bazylinski et al. 1988), and (3) bullet- or toothshaped (also described as anisotropic meaning these crystals lack a center of inversion symmetry; Mann et al. 1987a, b; Thornhill et al. 1994; Lefe`vre et al. 2011c). 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 crystal morphologies (Matsuda et al. 1983; Mann et al. 1984a, b, 1987a, b; Meldrum et al. 1993a, b). In crystallographic notation, numbers in square brackets (e.g., [100]) denote a direction vector. Coordinates in angle brackets (also referred to as chevrons), such as , denote a family of directions related by the symmetry of the crystal structure. The family of directions is called directions of a form. For cubic crystal structures, comprises eight directions (all the possible combinations of 1 and -1 taken three at a time). Numbers in parentheses such as (111) denote a particular plane of the crystal structure; the numbers are referred to as the Miller indices. Indices in curly brackets, such as {100}, represent a family of symmetry-related planes similar to the way angle brackets denote a family of directions (Borchardt-Ott 2011). Magnetotactic bacterial morphologies are derived from combinations of {111}, {110}, and {100} forms with suitable distortions (Devouard et al. 1998). The roughly cuboidal crystals are cuboctahedra ({100} + {111}), and the elongated, parallelepipedal crystals are either pseudo-hexahedral or pseudo-octahedral prisms derived from {100} + {110} + {111}. Examples are shown in > Fig. 12.17a–d. The cuboctahedral crystal morphology preserves the symmetry of the face-centered cubic spinel structure in which all equivalent crystal faces develop equally. The pseudo-hexahedral and pseudo-octahedral prismatic particles represent anisotropic growth in which equivalent faces develop unequally (Mann and Frankel 1989; Devouard et al. 1998). Synthesis of the bullet- and tooth-shaped magnetite particles (> Fig. 12.17e–h), the most anisotropic of the magnetotactic bacterial magnetite crystals, appears to be more complex than that of the other types. These crystals can be further subdivided into those with one pointed end and one flat end (flat-top shape; fts) and those with two pointed ends (double-triangular shape; dts) which appear as two isosceles triangles sharing a common base (> Fig. 12.18) (Lefe`vre et al. 2011c). In the dts crystals, both projected triangles appear to have the same width, although one triangle is longer than the other in mature crystals. The first of the anisotropic magnetosome crystals to be examined were fts type from an uncultured magnetotactic bacterium collected from the Exeter River in New Hampshire, USA. It was proposed that growth of these crystals occurred in two stages and that they have an idealized, six-sided prismatic, magnetosome habit comprising four {111} and two {100} faces capped by two faces of {111} with associated {111} and {100} corner faces. Crystal growth of a nascent cuboctahedron

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b

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. Fig. 12.16 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) cuboctahedrons; (b), (c), and (f) variations of elongated pseudohexagonal prisms; (d) elongated cuboctahedron. 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))

presumably commences via nucleation on the magnetosome membrane and continues until the width of the crystal is about 40 nm. In this first stage, there is proportional growth between width and length of the crystal. In the second stage, anisotropic growth commences with subsequent elongation parallel to while the crystal width remains relatively constant (Mann et al. 1987a, b). In magnetite magnetosome crystals with elongatedprismatic habits, the axis of elongation is the axis

of orientation which is considered the ‘‘easy’’ (lowest energy) direction of magnetization in single magnetite crystals (Frankel et al. 2007). When these particles are in a chain within a magnetotactic bacterial cell, they are oriented with the long crystal axis parallel to the chain axis. While elongated-anisotropic magnetosomes are also usually oriented with their long axes parallel to the chain axis, the axis of elongation varies among the , , , and axes (Lefe`vre et al. 2011c).

Magnetotactic Bacteria

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. Fig. 12.17 Brightfield TEM images of magnetite (Fe3O4) crystals in magnetosomes of different magnetotactic bacteria. (a) and (b), cuboctahedral crystals in the cultured gammaproteobacterium strain BW-2 isolated from Badwater Basin, Death Valley National Park, California (Lefe`vre et al. 2012); and from a spirillum, strain CB-1, isolated from Lake Mead, Nevada, respectively. (c) and (d), elongated-prismatic crystals from an uncultured magnetotactic bacterium found in sediment of the Mediterranean Sea collected at Marseille, France; and from the cultured, gammaproteobacterium strain SS-5 isolated from the Salton Sea, California (Lefe`vre et al. 2012), respectively. (e) through (h), elongated-anisotropic (bullet-shaped) crystals from a variety of different magnetotactic bacteria. (e) and (f), an uncultured, multicellular magnetotactic prokaryote (MMP) and an uncultured rod-shaped magnetotactic bacterium both found in sediment of the Mediterranean Sea collected at Marseille, France (Lefe`vre et al. 2007), respectively. (g), the uncultured, moderately thermophilic vibrio, Candidatus Thermomagnetovibrio paiutensis strain HSMV-1 (Lefe`vre et al. 2010b). (h), the uncultured ovoid magnetotactic bacterium, Ca. Magnetoovum mohavensis strain LO-1 found in sediment of Lake Mead, Nevada (Lefe`vre et al. 2011a)

There is evidence that in some cultured alkaliphilic magnetotactic bacteria, individual anisotropic crystals may partially result from aggregation of multiple magnetite crystals perhaps arising from multiple nucleation events in the magnetosome membrane vesicle (Lefe`vre et al. 2011c). Whereas the cuboctahedral form of magnetite can occur in inorganically formed magnetites (Palache et al. 1944), the prevalence of elongated-prismatic and elongated-anisotropic habits in magnetosome crystals imply anisotropic growth conditions, for example, 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 1989; Chang et al. 1989; Stolz et al. 1986, 1990; Stolz 1993). Iron Sulfide-Type Magnetosomes. Almost all known freshwater magnetotactic bacteria biomineralize magnetite as the mineral phase of their magnetosomes. In contrast, others, particularly many marine, estuarine, and salt marsh species, produce iron sulfide-type magnetosomes consisting primarily of the magnetic iron sulfide, greigite, and Fe3S4 (Heywood et al. 1990, 1991; Mann et al. 1990b; Po´sfai et al. 1998a, b) although these organisms have recently been found in nonmarine environments (Lefe`vre et al. 2011d). Early reports of nonmagnetic iron pyrite (FeS2; Mann et al. 1990b) and magnetic pyrrhotite (Fe7S8; Farina et al. 1990) have never been confirmed and likely represent misidentifications of additional iron sulfide species

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. Fig. 12.18 High magnification brightfield TEM images of elongatedanisotropic magnetite crystals in magnetotactic bacteria. These type of crystals can divided into those with one pointed end and one flat end (flat-top shape; fts) (a) and those with two pointed ends (double-triangular shape; dts) which appear as two isosceles triangles sharing a common base (b) (Lefe`vre et al. 2011c). In the dts crystals, both projected triangles appear to have the same width, although one triangle is longer than the other in mature crystals. The crystals in (a) are from the uncultured, moderate thermophilic magnetotactic bacterium, Candidatus Thermomagnetovibrio paiutensis strain HSMV-1 (Lefe`vre et al. 2010b) and those in (b) from a cell collected from sediment of an alkaline spring in California, of the now cultured obligately alkaliphilic magnetotactic bacterium strain AV-1 (Lefe`vre et al. 2011b)

occasionally observed with greigite in cells (Po´sfai et al. 1998a, b). Currently recognized greigite-producing magnetotactic bacteria includes the MMP (Farina et al. 1983; Rodgers et al. 1990a, b; DeLong et al. 1993) and a variety of relatively large, rod-shaped bacteria (Bazylinski et al. 1990, 1993a, b; Heywood et al. 1990, 1991; Bazylinski and Frankel 1992; Lefe`vre et al. 2011d). The iron sulfide-type magnetosomes contain either particles of greigite (Heywood et al. 1990, 1991) or a mixture of greigite and transient nonmagnetic iron sulfide phases that appear to represent mineral precursors to greigite (Po´sfai et al. 1998a, b). These phases include mackinawite (tetragonal FeS) and a possible sphalerite-type cubic FeS (Po´sfai et al. 1998a, b). Based on TEM observations, electron diffraction, and known iron sulfide chemistry (Berner 1967, 1970, 1974), the reaction scheme for greigite formation in the magnetotactic bacteria appears to be: cubic FeS ! mackinawite (tetragonal FeS) ! greigite (Fe3S4) (Po´sfai et al. 1998a, b). The de novo synthesis of nonmagnetic 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 might be expected to transform into pyrite (Berner 1967, 1970). However, pyrite has never been unequivocally identified in any magnetotactic bacterium. The same general morphologies of magnetite crystals in magnetotactic bacteria are also those observed for greigite (> Fig. 12.19): (1) cuboctahedral (the equilibrium form of face-centered cubic greigite) (Heywood et al. 1990, 1991), (2) elongated-prismatic (> Fig. 12.16f) (Heywood et al. 1990, 1991), and (3) bullet- and tooth-shaped (Po´sfai et al. 1998a, b). Like that of their magnetite counterparts, morphology of the greigite particles appears to be species- and/or strain-specific, although confirmation of this observation requires controlled studies of pure cultures of greigite-producing magnetotactic bacteria. One clear exception to this rule is the MMP (Farina et al. 1983; Bazylinski et al. 1990, 1993a; Mann et al. 1990b; Rodgers et al. 1990a, b; Bazylinski and Frankel 1992). This unusual microorganism contains pleomorphic, elongated-prismatic, bullet-shaped, and cuboctahedral greigite particles. Some of these particle morphologies are shown in > Figs. 12.4 and > 12.19c. Therefore, the biomineralization process appears to be more complicated in this organism than in the rods with greigite-containing magnetosomes or in magnetite-producing, magnetotactic bacteria.

Arrangement of Magnetosomes Within Cells of Magnetotactic Bacteria In cells of most magnetotactic bacteria, the magnetosomes are usually positioned as one or more chains that traverse the long axis of the cell (Bazylinski 1995; Bazylinski and Moskowitz 1997; Frankel and Moskowitz 2003) (> Figs. 12.17 and > 12.18). In the chain arrangement of the single magnetic domain crystal magnetosomes, the magnetic dipole moment of the cell is maximized because magnetic interactions between the magnetosomes cause each magnetosome moment to spontaneously orient parallel to the others along the chain axis by minimizing the magnetostatic energy (Frankel 1984; Frankel and Moskowitz 2003). Therefore, the total magnetic dipole moment of the chain and the cell is the algebraic sum of the moments of the individual crystals in the chain. This has been confirmed repeatedly using a number of techniques including direct magnetic measurements (Penninga et al. 1995), magnetic force microscopy (Proksch et al. 1995; Suzuki et al. 1998), and electron holography (Dunin-Borkowski et al. 1998, 2001). The significance of this is that the chain of magnetosomes in a magnetotactic bacterium functions like a single magnetic dipole rather than a collection of individual dipoles and causes the cell to behave similarly. Magnetotaxis results from this magnetic dipole imparted by the chain of magnetosomes which cause the cell to passively align along geomagnetic field lines while it swims (Frankel 1984; Frankel and Moskowitz 2003). Living cells are neither attracted nor pulled toward either geomagnetic pole, and dead cells, like living cells, also align along

Magnetotactic Bacteria

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control, assembly of magnetosome chains involves magnetostatic interaction, and magnetic ‘‘docking’’ to stable magnetic single domain particles is a key mechanism for building the functional cellular magnetic dipole (Faivre et al. 2010).

Biomineralization of Magnetosomes

. Fig. 12.19 Morphologies of intracellular greigite (Fe3S4) particles produced by magnetotactic bacteria. (a) Brightfield scanning transmission electron microscope (STEM) image of cuboctahedra in an unidentified rod-shaped 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 multicellular magnetotactic prokaryote (MMP) (Courtesy of M. Po´sfai and P. R. Buseck)

geomagnetic field lines but do not swim. Magnetosomes must be anchored in place within the cell to function as described as if they were free-floating in the cell, they would likely clump, causing a significant reduction in the cellular dipole moment. This is accomplished by dedicated cytoskeletal structures and close attachment to the inner cell membrane (see later section on > ‘‘The Magnetosome Membrane’’). In addition to biological

Because little is known regarding the biomineralization of greigite magnetosomes at the molecular level except that there is evidence that similar genes and proteins are involved (Abreu et al. 2011; Lefe`vre et al. 2011d), this section is focused on magnetite magnetosome synthesis. Biomineralization of the bacterial magnetosome appears to be a complex process that involves several steps that temporally overlap during the lifetime of the cell (> Fig. 12.20). The first step is invagination of the cell membrane and the possible formation of a bona fide, pinched off magnetosome membrane vesicle, an important question that remains unresolved. Using electron cryotomography, the magnetosome membrane in Magnetospirillum species has clearly been shown to originate as an invagination of the cytoplasmic membrane (CM) and that magnetite precipitation occurs after the invagination is formed (Komeili et al. 2006; Katzmann et al. 2010). Presumably, there is some sorting of magnetosome membrane proteins during the invagination and/or membrane vesicle formation process (Murat et al. 2010) as it is clear that magnetosome membranes contain proteins that are not present in the CM. Different stages of magnetite precipitation were observed within magnetosome membrane invaginations/ vesicles. Cells grown under iron limitation contained empty magnetosome invaginations/vesicles arranged in a chain engaged to the CM (Komeili et al. 2006). Only 35% of the magnetosomes examined showed the magnetosome membrane to be an invagination of the CM suggesting that the invaginations pinch off and become true vesicles. Alternatively, this may be a result of a technical problem involving the technique (Komeili 2007a, b). It is also not known if this is a common characteristic of magnetite magnetosomes in all magnetotactic bacteria. In parallel experiments with M. gryphiswaldense, Scheffel et al. (2006) found empty magnetosome membrane vesicles in cells grown under iron limitation and also that magnetic cells contain, in addition to magnetite filled magnetosome vesicles, many empty vesicles inside the cell. As in M. magneticum, vesicles in M. gryphiswaldense were shown by cryo-electron microscopy to invaginate from the CM (Katzmann et al. 2010). However, most mature vesicles appeared to be no longer connected to the CM, and it was therefore hypothesized that nascent magnetosome particles become detached during maturation of magnetite crystals in this organism (Faivre et al. 2007). The mature magnetosome membrane invaginations/vesicles probably become aligned in the chain motif during their formation. Iron uptake by the cell is certainly required for magnetosome synthesis and is likely occurring continually as long as it is available. Cells of cultured magnetotactic bacteria are extremely proficient at iron uptake as they have been shown to consist of

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. Fig. 12.20 (a) Cryo-electron tomogram of a section of cell of Magnetospirillum gryphiswaldense showing its intracellular organization. Magnetite crystals (orange) are closely adjacent to and aligned along bundles of the cytoskeletal magnetosome filament (yellow) formed by the actin-like MamK protein. Several vesicles of the magnetosome membrane (green) are visible. Blue represents outer and inner membranes. Figure modified from Katzmann et al. (2010). (b) Schematic representation of intracellular magnetosome formation in a magnetotactic bacterial cell. Extracellular iron (as ferrous or ferric ions) is taken up and transported into the cell. Biomineralization of magnetite crystals then occurs in specific compartments provided by the magnetosome membrane, in which the physico-chemical conditions required for magnetite crystallization are controlled. Magnetosome membrane vesicles originate by invagination from the inner membrane prior to magnetite synthesis. Mature magnetosome crystals are then assembled and concatenated into linear chains by the action of the magnetosome filaments, facilitated by magnetostatic interactions between particles

greater than 3 % iron on a dry weight basis, a value several orders of magnitude over non-magnetotactic bacterial species (Blakemore 1982; Heyen and Schu¨ler 2003). In addition, iron uptake for magnetite synthesis appears to occur relatively quickly (Schu¨ler and Baeuerlein 1998; Heyen and Schu¨ler 2003). It appears that both Fe(II) and Fe(III) can be taken up by cells of magnetotactic bacteria for magnetite synthesis although not necessarily simultaneously (Matsunaga and Arakaki 2007; Schu¨ler and Baeuerlein 1996; Suzuki et al. 2006). How iron is taken up by magnetotactic bacteria is unknown but it would seem that there would be multiple mechanisms for this in a single bacterium as has been found in other non-magnetotactic bacteria. Thus far, siderophores, low molecular weight ligands produced by the cell that chelate and solubilize Fe(III) (Neilands 1984, 1995), have been implicated

in iron uptake by magnetotactic bacteria (Paoletti and Blakemore 1986; Calugay et al. 2003; Dubbels et al. 2004) as well as a putative copper-dependent iron uptake system similar to that found in the yeast Saccharomyces cerevisae (Dubbels et al. 2004). By correlation of iron uptake rates with results from TEM analysis, it has been demonstrated that the morphology of magnetite crystals is not only determined by biological control through biological regulation at the magnetosome compartments but to some degree also by the rates of iron uptake by magnetotactic bacteria (Faivre et al. 2008). These observations imply that the biological control over magnetite biomineralization can be disturbed by environmental parameters. Iron then would have to enter the magnetosome invagination/vesicle. If the magnetite crystals are truly formed in permanent invaginations of the CM, then iron would only have to be transported through the outer membrane (OM) and enter the periplasm since any invagination of the CM would be open to the periplasm. This situation might only be temporary, however, if true independent vesicles are formed. In this case, iron may have to be transported across the CM and then through the magnetosome membrane to enter the vesicle. Several magnetosome membrane proteins have been implicated in this process (discussed in a later section). Based on Mo¨ßbauer spectroscopic analysis of Magnetospirillum gryphiswaldense, a mechanism was proposed by which iron required for magnetite biomineralization is processed throughout the CM directly to the magnetosome membrane without iron transport through the cytoplasm, suggesting that pathways for magnetite formation and biochemical iron uptake are distinct (Faivre et al. 2007). Magnetite formation occurs via membrane-associated crystallites, whereas the final step of magnetite crystal growth is possibly spatially separated from the CM. This work also suggests that cellular iron pools required for biochemical synthesis and magnetite biomineralization are distinct. This latter suggestion has been further substantiated by the analysis of a M. gryphiswaldense strain, in which the gene for a Fur-like iron uptake regulator was deleted (Uebe et al. 2010). This revealed that Fur is involved in global iron homeostasis, probably by balancing the competing demands for biochemical iron supply and magnetite biomineralization. In a very similar study, Qi et al. (2012) confirmed that Fur in M. gryphiswaldense directly regulates genes involved in iron and oxygen metabolism thereby influencing magnetosome biomineralization. Finally, there is nucleation and controlled maturation of the magnetite crystal within the magnetosome invagination/vesicle. Magnetite precipitation might occur through the reduction of hydrated ferric oxide(s) (Frankel et al. 1979, 1983; Schu¨ler and Baeuerlein 1998). However, when cells of Magnetospirillum gryphiswaldense were shifted from iron-limited to iron-sufficient conditions, they showed no delay in magnetite production (Heyen and Schu¨ler 2003) suggesting that there are no mineral precursors to magnetite during biomineralization or that they are unstable and convert to magnetite extremely quickly. The specificity for iron into the magnetosome mineral crystal appears to be very high. However, there are a number of

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. Fig. 12.21 (a) Purified magnetosomes of Candidatus Magnetococcus marinus negatively stained with uranyl acetate. Magnetosome membranes are represented by the electron-lucent layer surrounding each crystal. Note presence of chains. (b) After treatment with 1 % sodium dodecyl sulfate (SDS). Note electron-lucent layer on crystals is no longer present and the absence of chains

reports of the presence of other transition metals ions in magnetite and greigite magnetosome crystals in both cultured and uncultured magnetotactic bacteria. Trace amounts of titanium were found in magnetite particles of an uncultured freshwater magnetotactic coccus collected from a wastewater treatment pond (Towe and Moench 1981). The incorporation of small amounts of cobalt in surface layers of magnetosome magnetite crystals was demonstrated in three Magnetospirillum species (Staniland et al. 2008). Cells grown in cobalt-containing media showed very small changes in their magnetic properties, including the Verwey transition compared to a control culture. These results indicate that cobalt was not incorporated in the lattice structure of the magnetite crystals (Staniland et al. 2008). Uncultured magnetotactic bacteria in microcosms were exposed to MnCl2, and up to 2.8 % atomic manganese in ultrathin sectioned cells and magnetosomes was detected via localized energy dispersive X-ray analysis (Keim et al. 2009). Magnetic properties of these cells and their magnetosomes were not examined. Elemental maps of thin sections of magnetite magnetosomes showed a higher concentration of manganese at the edges of the crystals suggesting that, like cobalt in the previous study, manganese incorporation was limited to the surface of the crystals. Significant amounts of copper were found in greigite magnetosome crystals of some uncultured MMPs collected from a salt marsh in California (Bazylinski et al. 1993a). The concentration of copper was extremely variable and ranged from about 0.1 at.% to 10 at.% relative to iron. Again, copper appeared to be mostly concentrated on the surface of the crystals.

The Magnetosome Membrane The magnetosome membrane which encloses magnetite crystals (Gorby et al. 1988; Schu¨ler and Baeuerlein 1997) in magnetosomes appears to be the locus of control and regulation of magnetite biomineralization in magnetotactic bacteria

(Schu¨ler 2008) (> Fig. 12.21). It appears to be a universal feature of magnetotactic bacteria, although it has been speculated that at least one species, Desulfovibrio magneticus, may not possess magnetosome membranes around their magnetite crystals because of the inability to visualize them by several electron microscopic techniques (Byrne et al. 2010). Interestingly, despite this speculation, there has been a proteomic study of magnetosomes of D. magneticus to determine magnetosome membrane-associated proteins (Matsunaga et al. 2009). The magnetosome membranes of Magnetospirillum magnetotacticum and M. gryphiswaldense are lipid bilayers consisting of components typical of this type of membrane including proteins, fatty acids, glycolipids, sulfolipids, and phospholipids (Gorby et al. 1988; Gru¨nberg et al. 2004). This is in contrast to other intracellular inclusions in prokaryotes which are generally surrounded by a single layer of protein. Phospholipids make up 58–65 % of the total lipids of the magnetosome membrane of M. magneticum (Nakamura and Matsunaga 1993), and the predominant phospholipids in all Magnetospirillum species are phosphatidylserine, phosphatidylglycerol, and phosphatidylethanolamine (Gorby et al. 1988; Nakamura and Matsunaga 1993; Gru¨nberg et al. 2004). A comparison of the fatty acids of the magnetosome membrane, the CM and the outer membrane (OM) showed that the composition of the magnetosome membrane is similar to the CM but distinct from the OM (Tanaka et al. 2006) suggesting that the magnetosome membrane is derived from the CM. In addition, magnetite magnetosomes are almost always located adjacent to the CM in Magnetospirillum species (Bazylinski and Schu¨bbe 2007; Katzmann et al. 2010).

Magnetosome Membrane Proteins Magnetosome membranes can be easily removed from magnetosomes using detergents such as sodium deodecyl sulfate

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(SDS) for protein analysis (> Fig. 12.21). Protein profiles of the magnetosome membrane are distinct from other cell fractions (the soluble periplasmic and cytoplasmic fractions, and the cell and outer membrane fractions) in currently recognized Magnetospirillum species, in Desulfovibrio magneticus, and in Magnetovibrio blakemorei (Gorby et al. 1988; Okamura et al. 2000; Gru¨nberg et al. 2001; Dubbels et al. 2004; Tanaka et al. 2006; Matsunaga et al. 2009). In addition, there are also differences in the magnetosome membrane protein profiles between these organisms and even between the species of Magnetospirillum (Gru¨nberg et al. 2004). Because the magnetosome membrane contains proteins that are unique to it, it seems very probable that these proteins play the key roles in magnetite biomineralization in magnetosomes. Most of the focus of investigators in magnetite biomineralization by magnetotactic bacteria is on these proteins and the genes that encode for them. These proteins and genes are generally referred to as the Mam (magnetosome membrane) or Mms (magnetic particle membrane specific) proteins and the mam or mms genes, respectively, although a gene called magA has been described as coding for a magnetosome membrane protein (Matsunaga et al. 1992; Nakamura et al. 1995b). All mam and mms genes have been found to be clustered within several operons of the conserved genomic magnetosome island (see below). Identifying the function of the magnetosome membrane proteins appears to be key to understanding magnetosome biomineralization. Putative functions of these proteins, based on comparisons of similar proteins through blast searches and through mutagenesis experiments, include iron uptake into the cell and/or the magnetosome vesicles, crystal nucleation and biomineralization of magnetite, and the arrangement of the magnetosomes in the chain motif. The putative roles of a number of magnetosome membrane proteins follow, although this list is not complete. It should also be kept in mind that each magnetotactic bacterium appears to have genes within their magnetosome gene islands that are specific to them that might encode for proteins involved in biomineralization that have little or no homology with any known proteins. The MagA protein of Magnetospirillum magneticum has low similarity to the Escherichia coli potassium efflux protein KefC and its transcription reported to be upregulated by low iron concentrations in the growth medium (Nakamura et al. 1995b). Based on the putative phenotype of a non-complemented transposon mutant, a potential function is as a magnetosomedirected ferrous iron transporter having an essential role in magnetosome formation in M. magneticum (Nakamura et al. 1995a). However, a recent study showed that targeted deletion magA mutants of M. magneticum and M. gryphiswaldense still biomineralize wild type-like magnetosomes and have no obvious growth defects, thus unambiguously showing that magA is not involved in magnetosome formation in magnetotactic bacteria (Uebe et al. 2012). Based on genetic studies in Magnetospirillum magneticum, four genes (mamI, mamL, mamQ, and mamB) seem to be absolutely essential for the formation of magnetosomes (Murat et al. 2010) but were not sufficient to support

magnetosome formation in the absence of other magnetosome genes. MamI and MamL are unique to magnetotactic bacteria and were implicated in the invagination of the magnetosome membrane from the CM, since in DmamI and DmamL mutants, no structures resembling empty magnetosome compartments could be detected. However, the mechanism by which this is mediated is unclear, and MamI and MamL lack any significant homology to eukaryotic proteins known to be involved in deformation of cellular membranes. The mamA (corresponds to mam22 and mms24 in different magnetotactic bacteria) gene is present in the genomes of all magnetotactic bacteria examined (Okuda et al. 1996; Gru¨nberg et al. 2001; Komeili et al. 2004; Matsunaga et al. 2005; Schu¨bbe et al. 2009; Nakazawa et al. 2009; Abreu et al. 2011). The amino acid sequences of the MamA proteins show high similarity to proteins of the tetratricopeptide repeat (TPR) protein family (Okuda et al. 1996). MamA is thought to be important in protein-protein interactions that might occur in the synthesis of magnetosomes and the magnetosome chain (Okuda et al. 1996; Okuda and Fukumori 2001) since multiple copies of TPRs are known to form scaffolds within proteins to mediate protein-protein interactions and to coordinate the assembly of proteins into multisubunit complexes (Ponting and Phillips 1996). A deletion of mamA in Magnetospirillum magneticum resulted in shorter magnetosome chains, this leading to the suggestion that MamA activates magnetosome vesicles and is involved in magnetite crystal maturation (Komeili et al. 2004; Murat et al. 2010). Genes for the proteins MamB and MamM are also present in the genomes of all magnetotactic bacteria examined (Gru¨nberg et al. 2001; Matsunaga et al. 2005; Schu¨bbe et al. 2009; Nakazawa et al. 2009; Abreu et al. 2011) and show strong similarity to heavy metal transporting proteins of the cation diffusion facilitator family. An additional magnetosome membrane protein, MamV, also appears to be in this family, but its gene is only present in M. magnetotacticum and M. magneticum and not in other magnetotactic bacteria. Proteins in this family display an unusual degree of size variation, sequence divergence, and polarity, can catalyze the influx or efflux of metal ions, and include a ferrous iron transport system (Paulsen et al. 1997; Grass et al. 2005; Haney et al. 2005). For this reason, these Mam proteins might be involved in the transportation of the iron into the magnetosome vesicle (Gru¨nberg et al. 2001). As demonstrated by a recent study in M. gryphiswaldense, MamB and MamM form heterodimers and also interact with other magnetosome proteins, indicating that the functions of these two proteins are complex and involved in the control of different key steps of magnetosome formation (Uebe et al. 2012). Genes for the MamE, MamO, and MamP proteins are present in all magnetotactic bacteria investigated to date. MamE is required for magnetite biomineralization in Magnetospirillum magneticum (Murat et al. 2010). These proteins show sequence similarity to HtrA-like serine proteases but little similarity to each other. HtrA (also known as DegP) is a heat-shock-induced, envelope-associated serine protease first discovered in Escherichia coli (Lipinska et al. 1989). The main role of HtrA

Magnetotactic Bacteria

seems to be in the degradation of misfolded proteins in the periplasm (Pallen and Wren 1997). These proteases are also known to be involved in nondestructive protein processing and modulation of signaling pathways by degrading important regulatory proteins and are characterized by the inclusion of one or two PDZ-domains (Fanning and Anderson 1996) and a trypsin-like protease domain. These proteins could function as chaperones in magnetosome formation (Gru¨nberg et al. 2001). In M. magneticum, MamE may be important in arranging proteins in the magnetosome membrane while MamO may be involved in iron uptake and magnetosome magnetite crystal initiation in the magnetosome invagination (Murat et al. 2010). Both MamE and MamO have been shown to be essential for magnetite biomineralization in M. gryphiswaldense based on results from gene deletion experiments (Yang et al. 2010). The magnetosome membrane proteins MamC (Mms13, Mam12 (Arakaki et al. 2003; Taoka et al. 2006)), MamD (Mms7; Fukuda et al. 2006), MamF, MamG, MamQ, MamR, and MamS are unique to some magnetotactic bacteria, and homologues of these proteins have not been found in nonmagnetotactic bacteria (Gru¨nberg et al. 2004). All recognized Magnetospirillum species contain the genes for these proteins. Candidatus Magnetococcus marinus contains all but mamG and mamR (Schu¨bbe et al. 2009). Magnetovibrio blakemorei contains all but mamG in its magnetosome gene island, but the presence of the other genes in the genome cannot be excluded at this time (genome of this organism is not complete; Jogler et al. 2009a). Desulfovibrio magneticus possesses only mamQ (Nakazawa et al. 2009). Ca. Magnetoglobus multicellularis and Ca. Magnetobacterium bavaricum contain only mamQ, the latter species has two copies, but the presence of the other genes cannot be excluded as the genomes are not complete Abreu et al. 2011; Jogler et al. 2011). MamC is an abundant protein in the magnetosome membranes of M. magnetotacticum (Taoka et al. 2006), M. gryphiswaldense (Gru¨nberg et al. 2001), and Magnetovibrio blakemorei (Dubbels et al. 2004). The hydrophobic proteins, MamC and MamF, contain predicted transmembrane helices. MamD and MamG are partially identical and are similar to another magnetosome membrane protein, Mms6 of M. magneticum. All three proteins contain large repeating leucine-glycine (L-G) motifs present in other proteins known to be involved in biomineralization. Mms6, an amphiphilic protein consisting of an N-terminal LG-rich hydrophobic region and a C-terminal hydrophilic region containing repeats of acidic amino acids, has been shown to affect the crystal morphology of crystals when present during the chemical precipitation of magnetite (Arakaki et al. 2003; Prozorov et al. 2007). The proteins MamGFDC, in concert, comprise about 35% of the protein associated with the magnetosome membrane and, although not essential for magnetite biomineralization, have been shown to regulate the size of magnetosome magnetite crystals in M. gryphiswaldense (Scheffel et al. 2008). In M. magneticum, MamR and MamS appear to be involved in magnetite crystal maturation while MamQ in the invagination of the cytoplasmic membrane to form the magnetosome vesicle (Murat et al. 2010; Quinlan et al. 2011).

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MamN, the gene of which is not present in the genomes of Candidatus Magnetococcus marinus and Desulfovibrio magneticus (Schu¨bbe et al. 2009; Nakazawa et al. 2009) and also not in the putative magnetosome gene islands of Ca. Magnetoglobus multicellularis and Ca. Magnetobacterium bavaricum (Abreu et al. 2011; Jogler et al. 2011), shows some similarity to certain transport proteins, some of which transport protons leading to an idea that this protein might function as a proton pump transporting protons accumulating during magnetite precipitation (Jogler and Schu¨ler 2007). In Magnetospirillum magneticum, MamN, like MamM, is thought to be involved in iron uptake and initiation of magnetite crystal formation (Murat et al. 2010). The gene for MamT is present in all magnetotactic bacteria studied (Gru¨nberg et al. 2001; Matsunaga et al. 2005; Schu¨bbe et al. 2009; Nakazawa et al. 2009; Abreu et al. 2011) except in the putative magnetosome gene island of Candidatus Magnetobacterium bavaricum (Jogler et al. 2011). MamT contains two possible binding sites for the heme group present in cytochrome c and, therefore, might be involved in redox reactions within the magnetosome vesicle (Gru¨nberg et al. 2004) that might be important in magnetite crystal maturation (Murat et al. 2010). The genes mamJ and mamK are located within the mamAB gene cluster in Magnetospirillum species and are cotranscribed (Schu¨bbe et al. 2006). The mamK gene has been found in all magnetotactic bacteria studied except in the putative magnetosome gene island of Candidatus Magnetobacterium bavaricum (Jogler et al. 2011), while mamJ is present only in recognized Magnetospirillum species (Nakazawa et al. 2009; Schu¨bbe et al. 2009; Jogler et al. 2009a, 2011; Abreu et al. 2011). MamJ is a strongly acidic protein with a repeating glutamate-rich section in its central domain (Scheffel et al. 2006) that is reminiscent to certain other acidic proteins (Gru¨nberg et al. 2004) involved in biomineralization processes such as calcium carbonate biomineralization in shells (Baeuerlein 2003). Carboxy groups of the acidic amino acids are recognized to have a high affinity for metal ions, and because of this, magnetosome proteins with these characteristics have been thought to be involved in the initiation of magnetite crystal nucleation (Arakaki et al. 2003). However, deletion of mamJ in M. gryphiswaldense had no effect on the biomineralization of magnetite but resulted in cells in which magnetosome crystals were organized in agglomerated clusters instead of regular chains (Scheffel et al. 2006), whereas in M. magneticum, the phenotype of a co-deletion of mamJ along with the paralogous limJ gene was less severe and resulted in interrupted magnetosome chains (Draper et al. 2011). MamK shows some homology to actin-like proteins including MreB (Schu¨bbe et al. 2003), which have important functions in the control of cell morphology and elongation, peptidoglycan synthesis, and portioning of plasmids in many bacteria (Jones et al. 2001; Figge et al. 2004; Carballido-Lopez 2006; Cabeen and Jacobs-Wagner 2010; Garner et al. 2011; Dominguez-Escobar et al. 2011). MamK proteins in magnetotactic bacteria are more similar to each other than they are to MreB homologues (Komeili et al. 2006; Derman et al. 2009). In addition, assembly of

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MamK filaments appears to be independent of MreB (Pradel et al. 2006). Experiments involving gene knockout mutants of mamJ in M. gryphiswaldense and mamK in M. magneticum showed that the products of these genes are responsible for magnetosome chain formation but did not inhibit magnetosome formation in these organisms (Komeili et al. 2006; Scheffel et al. 2006). MamJ is thought to function by anchoring magnetosomes to MamK filaments in Magnetospirillum species (Komeili et al. 2006; Scheffel et al. 2006; Scheffel and Schu¨ler 2007), whereas MamK is involved in dynamic assembly, positioning and segregation of the magnetosome chain during cell cycle rather than merely providing a rigid mechanical scaffold (Katzmann et al. 2010, 2011). However, the observed differences between mutant phenotypes in different magnetospirilla suggest that the functions of mamK and mamJ may be somewhat distinct in different species depending on their genetic context.

Genomics and Genetics of Magnetotactic Bacteria The genome sequences of several magnetotactic bacteria are now complete or nearly so and are available for examination. Analysis of these genome sequences has provided valuable insight into how magnetosome genes are organized in different magnetotactic bacteria. The genome of Magnetospirillum magnetotacticum strain MS-1 consists of a single, circular chromosome about 4.3 Mb in size with a possible extrachromosomal element as determined by pulsed-field gel electrophoresis (Bertani et al. 2001). The genome of this bacterium is partially sequenced and annotated and is available at the Joint Genome Institute’s website (http://genome.jgi-psf.org/draft_microbes/ magma/magma.home.html). M. magneticum strain AMB-1 contains a circular chromosome slightly larger than that of M. magnetotacticum at 4.97 Mb (Matsunaga et al. 2005) and likely a cryptic plasmid (Okamura et al. 2003). The genome sequence of this species is available at the DNA Data Bank of Japan (http:// www.ddbj.nig.ac.jp) under accession number AP007255. The genome of M. gryphiswaldense strain MSR-1 is comprised of a circular chromosome about 4.3 Mb in size and also contains a native plasmid (Jogler and Schu¨ler 2007; Richter et al. 2007). The genome of the marine coccus Candidatus Magnetococcus marinus strain MC-1 consists of a singular, circular chromosome about 4.5 Mb in size and there is no evidence for the presence of plasmids. The genome sequence of this species is complete (Schu¨bbe et al. 2009) and can be viewed at http://genome.jgi-nsf.org/draft_microbes/magm1/magm1. home.html. The genomes of the marine vibrios Magnetovibrio blakemorei strains MV-1 and MV-2 are approximately 3.7 and 3.6 Mb in size, respectively, based on pulsed-field gel electrophoresis. Data suggest genomes of both strains consist of a single, circular chromosome with no evidence of plasmids (Dean and Bazylinski 1999). The genome of MV-1 is almost complete. The genome sequence for Desulfovibrio magneticus strain RS-1 is complete (Nakazawa et al. 2009) and is available

at the DNA Data Bank of Japan (http://www.ddbj.nig.ac.jp) under accession numbers AP010904 to AP010906. In an attempt to identify magnetotaxis-specific genes by bioinformatic analysis, cross-comparisons of the complete or nearly complete genomes of the magnetotactic Alphaproteobacteria including Magnetospirillum magneticum, M. magnetotacticum, M. gryphiswaldense, and Candidatus Magnetococcus marinus revealed a core genome of about 890 genes which are shared by all four strains. In addition to a set of approximately 152 genusspecific genes shared by the three Magnetospirillum strains, 28 genes were identified as group specific, that is, which occur in all four analyzed magnetotactic Alphaproteobacteria but exhibit no (magnetotactic bacterial-specific genes) or only weak (magnetotactic bacterial-related genes) similarity to any genes from non-magnetotactic organisms and which, besides various novel genes, included nearly all mam and mms genes that were previously shown to control magnetosome formation. If the genome sequence of the sulfate-reducing, deltaproteobacterium Desulfovibrio magneticus was available at the time for inclusion in this comparison, the number of signature genes conserved in all five magnetotactic Proteobacteria decreased to only nine.

Molecular Organization of Magnetosome Genes In all magnetotactic bacteria examined, the mam and mms genes are present as clusters that are in relatively close proximity to one another within the genome and are organized as a genomic ‘‘magnetosome island (MAI)’’ (see below). The mamA and mamB genes in Magnetospirillum gryphiswaldense are present in a segment of DNA about 16.4 kb in length in collinear order with 15 other genes that comprise the mamAB cluster (Gru¨nberg et al. 2001). Recent deletion studies in M. magneticum and M. gryphiswaldense demonstrated that the mamAB cluster is the only operon-containing genes that are absolutely essential for magnetosome formation and magnetite biomineralization, whereas the other operons have important accessory functions in controlling the synthesis of regularly shaped and sized crystals that are functional for magnetic orientation (Murat et al. 2010; Ullrich and Schu¨ler 2010; Lohße et al. 2011). One of such accessory operons, the mamGFDC cluster is about 2.1 kb in length located about 15-kb upstream of the mamAB operon and is composed of four genes which encode a group of abundant magnetosome membrane proteins involved in size control of magnetite crystals (Scheffel et al. 2008). The 3.6 kb mms6 cluster is located 368 bp upstream of the mamGFDC operon and contains five genes (Schu¨bbe et al. 2003). Another gene encoding for a magnetosome membrane protein, mamW, is not present in these three operons but is located about 10-kb upstream of the mms6 operon (Ullrich et al. 2005). All mam and mms genes are located on a segment of DNA about 45 kb in length in M. gryphiswaldense. The operon-like, collinear organization of the mamAB, mamGFDC, and mms6 clusters suggests that they are transcribed as single long mRNAs, and experimental evidence provides support for this polycistronic transcriptional unit. The

12

Magnetotactic Bacteria

Magnetospirillum gryphiswaldense mms6 G F D C

W

H

J

E

I

K

L

O

N

M

P

A

B like

U

S T

Q R B

Z

Y

X

Magnetospirillum magneticum C D F mms6

E

I

H

J

K

L

M

N

O

P

A

Q R B

S T

U

V

J

K

L

M

N

O

P

A

Q R B S T

U

V

K

F

W

O

E

Z

Q R B

X

Y

F

Magnetospirillum magnetotaticum C D

F

mms6

E

I

H

O

E

Z

Q R B

X

Y

F

W

Candidatus Magnetococcus marinus mms6

H

I

E

C

M

O

P

A

A like

Q

K

H

B

S T

D like

Z

X

D

A

Candidatus Magnetovibrio blakemorei I

E

K

M

N

O

P

A

Q R B

S T

Y

D like mms6

C

X

Z

R like

Desulfovibrio magneticus A

I

Q

B

T

P

E

O

M

K 1000 bp

Candidatus Magnetobacterium bavaricum P

M

Q

B

A I

E

Q

Candidatus Magnetoglobus multicellularis K

P

E

B

A

Q

M

P like

T

O

TPR protein

CDF transporter

Generic transporter

Acidic repetitive protein

Actin-like MreB protein

LemA-like protein

PDZ domain (HtrA-like serine protease) MTB-specific genes

Hypothetical protein General magnetosome gene

. Fig. 12.22 Organization of magnetosome genes in the putative magnetosome gene islands (MAIs) of different magnetotactic bacteria. All organisms are cultured with the exception of Candidatus Magnetobacterium bavaricum and Ca. Magnetoglobus multicellularis. Letters above genes indicate mam genes (e.g., A mamA)

transcription starting points of the mamAB, mamGFDC, and mms6 operons were mapped closely upstream of the first genes in the operons, respectively (Schu¨bbe et al. 2006). The organization of the mam and mms genes is relatively well conserved in Magnetospirillum strains. In addition, there are high similarities for specific Mam and Mms proteins and their encoding genes, respectively, in recognized Magnetospirillum species. The organization and sequence of the magnetosome genes is less conserved in other unrelated magnetotactic strains (Schu¨bbe et al. 2003, 2009; Ullrich et al. 2005; Jogler et al. 2011). The genomic region that contains the magnetosome genes in Magnetospirillum gryphiswaldense also contains 42 mobile elements as transposases of the insertion sequence type and integrases (Ullrich et al. 2005). These mobile elements are common, important features in genomic islands (Mahillon and Chandler 1998; Mahillon et al. 1999). Other characteristics of gene islands include the presence of tRNA genes that can act as insertion sites for integrases (Blum et al. 1994; Reiter and Palm 1990) and a different guanine + cytosine (G + C) content compared to the rest of the genome (Dobrindt et al. 2004). In M. gryphiswaldense, the magnetosome gene region is about 130 kb in size, contains three tRNA genes upstream of the mms operon, has a slightly different G + C content versus the rest of the genome, and contains many hypothetical genes and pseudogenes (Schu¨bbe et al. 2003; Ullrich et al. 2005) which apparently have no function as their deletions had no obvious effect on either growth or magnetosome formation

(Lohße et al. 2011). Therefore, it seems very likely that this genomic region represents a large magnetosome gene island (MAI) which appears to be present with variations in other cultured and uncultured magnetotactic bacteria (Fukuda et al. 2006; Richter et al. 2007; Nakazawa et al. 2009; Schu¨bbe et al. 2009; Abreu et al. 2011; Jogler et al. 2011). A comparison of the putative MAIs of different cultured and uncultured magnetotactic bacteria is shown in > Fig. 12.22. > Table 12.1 lists all the magnetosome genes present in the putative MAIs of all magnetotactic bacteria in which one has been identified. Gene or genomic islands are reported to be distributed to different bacteria through horizontal gene transfer and thus may be a major pathway for the evolution of bacterial genomes (Juhas et al. 2009). In addition, genomic islands are thought to undergo frequent gene rearrangements (Juhas et al. 2009). Gene rearrangements, gene deletions, and duplications may be the reason for the frequent development of spontaneous nonmagnetic mutants of various strains. Spontaneous deletions that lead to a loss of the magnetic phenotype with a frequency of 10 2 were observed under starvation conditions in late stationary phase cultures of Magnetospirillum gryphiswaldense and most likely caused by RecA-dependent homologous recombination between numerous repeats present in the MAI (Ullrich et al. 2005; Kolinko et al. 2011). One of these mutants, designated M. gryphiswaldense strain MSR-1B, showed poorer growth in the presence of increased iron concentration and lower iron uptake compared to the wild-type strain (Schu¨bbe et al. 2003). Frequent

481

482

12

Magnetotactic Bacteria

nonmagnetic mutants that do not synthesize magnetosomes were also observed in Magnetovibrio blakemorei (Dubbels et al. 2004) and M. magneticum (Fukuda et al. 2006; Komeili et al. 2006). Rioux et al. (2010) identified a separate group of mam-like genes in the genome of Magentospirillum magneticum strain AMB-1. These genes, including mamKDLJEFQ-like genes, are clustered in a genomic islet distinct and distant from the known

magnetosome genomic island. In this study, they demonstrate that the mamK-like gene is transcribed and that the gene product is protein filaments as is MamK. Thus far, this is the only report of a functional mam gene located outside of the magnetosome genomic island. There is also some evidence for the presence of magnetosome membrane protein genes on a plasmid rather than a genome in magnetotactic bacterium (Matsunaga et al. 2009). Genes encoding for two homologous

. Table 12.1 Magnetosome genes present in the putative magnetosome gene islands of different cultured and uncultured magnetotactic bacteria Gene

MSR-1a b

MS-1

mamA

+

+

mamB

+

+

c

e

AMB-1

MC-1

MV-1

RS-1

Ca. M. bav.

Ca. M. mult.

+

+

+

+

+

+

+

+

+

+

+

+

d

d

mamC

+

+

+

+

+

/

/

mamD

+

+

+d

++

+

/

/

mamE

+

+

+

+

+

+

+

mamF

+

+

+

+

mamG

+

+

+

mamH

+

+

+

+

mamI

+

+

+

+

mamJ

+

+

+

mamK

+

+

+

mamL

+

+

+

mamM

+

+

+

+

+

/

/

/

/

/

+

/

/

+

+

/ +

++

+

+ +

+

+

/ /

/

+

/

/

+

+

mamN

+

+

+

/

/

mamO

+

+

+

+

+

+

/

+

mamP

+

+

+

+

+

+

+

+

mamQ

+

+

+

+

+

+

++

+

mamR

+

+

+

mamS

+

+

+

+

mamT

+

+

+

+

/

+

mamU

+

+

+

/

/

/

mamV

–

+

+

/

/

/

mamW

+

mamX

+

+

+

+

+ /

+

+

/

/

+

/

/

+

+

/

/

/

+

+

/

/

+

/

/

+

+

/

/

/

/

/

mamY

+

+

+

mamZ

++

++

++

mgI462

+

+

+

mms6

+

+

+

+

+

/

/

mgI459

+

+

+

+

+

/

/

mgI458

+

+

+

/

/

/

mgI457

+

+

+

/

/

/

mamE/S-like

+

+

+

+

/

/

/

mamF-like

+

+

+

+

/

/

/

mamH-like

+

+

+

+

/

/

/

mamA-like

–

++

/

/

/

/

/

/

mamP-like

–

mgr4150

+

+ +

+

Magnetotactic Bacteria

12

. Table 12.1 (continued) Gene

MSR-1a

MS-1

AMB-1

MC-1

MV-1

Ca. M. bav.

Ca. M. mult.

mgr0208

+

+

+

+

/

/

/

mgr0207

+

+

+

+

/

/

/

mgr0206

+

+

+

+

/

/

/

mgr3500

+

+

+

+

/

/

/

/

/

/

/

/

/

/

/

/

mgr3499

+

+

+

mgr3497

+

+

+

mgr3495

+

+

+

RS-1

a

Organisms: MSR-1, Magnetospirillum gryphiswaldense (Ullrich et al. 2005); MS-1, M. magnetotacticum (Bertani et al. 2001); AMB-1, M. magneticum (Matsunaga et al. 2005); MC-1, Candidatus Magnetococcus marinus (Schu¨bbe et al. 2009); MV-1, Magnetovibrio blakemorei (Jogler et al. 2009a); RS-1, Desulfovibrio magneticus (Nakazawa et al. 2009); Ca. M. bav. Ca. Magnetobacterium bavaricum (Jogler et al. 2011); Ca. M. mult. Ca. Magnetoglobus multicellularis (Abreu et al. 2011) b Symbols: +, homologue present in genome; ++, two paralogues in genome; -, homologue absent from genome; /, homologue absent from putative magnetosome island but genome sequence has not been completed c In M. magnetotacticum strain MS-1, mamA = mam22 (Okuda et al. 1996) d In M. magneticum strain AMB-1, mamA = mms24, mamC = mms13 and mamD = mms7 (Fukuda et al. 2006) e In M. magnetotacticum strain MS-1, mamC = mam12 (Taoka et al. 2006)

magnetosome proteins of Candidatus Magnetococcus marinus were found on a cryptic plasmid (pDMC1) of Desulfovibrio magneticus (Matsunaga et al. 2009). Strain BW-1, the only greigite-producing magnetotactic bacterium known to be in pure culture, biomineralizes both greigite and magnetite and contains two sets of magnetosome genes although it is not known whether they are on separate gene islands (Lefe`vre et al. 2011d). Because one set of genes is more similar to the magnetite-producing Desulfovibrio magneticus and Candidatus Magnetococcus marinus and the other to the greigite-producing Candidatus Magnetoglobus multicellularis, it was suggested that the first set is responsible for greigite biomineralization and the second for magnetite production (Lefe`vre et al. 2011d). Because the proportion of the different minerals produced is affected by external conditions in the growth medium, perhaps the two sets of genes are regulated separately. Distribution of the MAI through horizontal gene transfer would explain the phylogenetic diversity of the magnetotactic bacteria, while variations of the MAI in different magnetotactic bacteria may be the result of rearrangements within the MAI occurring over time.

Genetic Manipulation of Magnetotactic Bacteria Because of the difficulty in growing magnetotactic bacteria on agar plates as individual colonies, it took many years to develop tractable genetic systems in these organisms. In general, they do not form colonies unless the agar plates contain compounds to scavenge toxic radicals (e.g., activated charcoal) or they are incubated under low concentrations of oxygen. The most common way of assigning definitive functions to specific genes in prokaryotes is through single gene knockouts with subsequent analysis of the mutant phenotype, and this approach has proved very useful in the determination of the functions of a number of

magnetosome genes. Tractable genetic systems have now been developed for Magnetospirillum gryphiswaldense strain MSR-1 (Schultheiss and Schu¨ler 2003) and M. magneticum strain AMB-1 (Matsunaga et al. 1992). It is relatively easy to detect non-magnetotactic mutants of magnetotactic bacteria unable to synthesize magnetosomes since magnetite-forming colonies are generally dark brown or black versus those of nonmagnetic mutants which are white or cream colored (Dubbels et al. 2004; Schultheiss and Schu¨ler 2003). In addition, the degree of a magnetic response or its absence related to the number of magnetosomes per cell can be easily tested by light-scattering measurements of cell suspensions in variable magnetic fields by ‘‘Cmag’’ values (Schu¨ler et al. 1995; Zhao et al. 2007). Conjugational transfers of replicative plasmids were accomplished with frequencies of 1 and 3 4.5  10 3 for Magnetospirillum gryphiswaldense and M. magneticum, respectively (Matsunaga et al. 1992; Schultheiss and Schu¨ler 2003). Mutants of both strains were generated using Tn5 transposon mutagenesis as well as broad host range replication (pBBRMCS, IncQ) and suicide vectors (pK19mobsacB, pMB1) (Komeili et al. 2004; Matsunaga et al. 1992; Schultheiss and Schu¨ler 2003; Schultheiss et al. 2004). The development of genetic systems for these strains has allowed for the extrachromosomal expression of genes and the integration of reporter genes like luciferase or green fluorescent protein genes (gfp) and their derivatives. In turn, these techniques have allowed for studies involving the subcellular localization of proteins putatively involved in magnetite magnetosome biomineralization (Komeili et al. 2004; Matsunaga et al. 2000a, b; Nakamura et al. 1995b; Schultheiss et al. 2004). General transposon mutagenesis is random but can generate nonmagnetic mutants that make it possible to identify genes in the genome involved in magnetite biomineralization. Suicide vectors together with genomic data now allow for the integration of these vectors at specific sites on the genome to generate site-directed gene knockouts for the definitive

483

484

12

Magnetotactic Bacteria

determination of precise roles of specific genes in magnetite magnetosome biomineralization (Komeili et al. 2004, 2006; Pradel et al. 2006; Scheffel et al. 2006; Murat et al. 2010). Related techniques based on the Cre-lox system have also allowed the targeted deletion of large (up to 60 kb and more) regions from the genome of M. gryphiswaldense, which facilitates functional analysis and genomic engineering (Ullrich and Schu¨ler 2010; Lohße et al. 2011).

Applications of Magnetotactic Bacteria, Magnetosomes, and Magnetosome Crystals Cells of magnetite-producing magnetotactic bacteria and their magnetic inclusions have been shown to have novel magnetic, physical, and optical properties that can and have been exploited in a variety of scientific, commercial, and other applications (reviewed in Lang and Schu¨ler 2006; Lang et al. 2007; Matsunaga and Arakaki 2007; Arakaki et al. 2008; Xie et al. 2009). While the number of applications and patents involving magnetotactic bacteria appears to be ever increasing, a major problem is the mass culture of these organisms and the subsequent efficient harvesting of magnetosomes. However, there has been significant progress in this area in the last decade.

Mass Culture of Magnetotactic Bacteria Considering that the amount of magnetic materials from magnetotactic bacteria required for most applications is relatively high, obtaining higher yields of magnetotactic bacterial cells and magnetosomes from cultures poses a significant obstacle. In order to produce enough cells, magnetosomes, and magnetite crystals for these applications, cells therefore must be grown in mass culture where conditions for growth and magnetite synthesis must be optimized. In almost all cases, the focus of these studies involved modification of growth media and the conditions under which cultures are incubated. Magnetospirillum species are the only magnetotactic bacteria used in these studies. In some cases, it is difficult to compare yields directly as some studies focus on magnetosomes and it is unclear whether magnetosome membranes are included in the yield values. In an early study of this type, Matsunaga et al. (1990) grew Magnetospirillum magneticum in a 1,000-l fermenter and recovered 2.6 mg of magnetosomes per liter of culture. Culture optimization experiments were later conducted in fed-batch cultures of the same organism but did not result in a higher yield of cells or magnetosomes (Matsunaga et al. 1996, 2000a). A recombinant M. magneticum strain harboring the plasmid pEML was grown in a pH-regulated fed-batch culture system where the addition of fresh nutrients was feedback-controlled as a function of the pH of the culture (Yang et al. 2001a). Here, the magnetosome yield was maximized by adjusting the rate of addition of the major iron source. Providing ferric quinate at 15.4 mg/min resulted in a magnetosome yield of 7.5 mg/l.

Different iron sources and the addition of various nutrients and chemical reducing agents (e.g., L-cysteine, yeast extract, polypeptone) were also later shown to have significant effects on magnetosome yield by M. magneticum grown in fed-batch culture (Yang et al. 2001b). More precise control over the growth of Magnetospirillum species was achieved using an oxygen-controlled fermenter (Heyen and Schu¨ler 2003; Lang and Schu¨ler 2006). Three species were grown using this method, M. gryphiswaldense, M. magnetotacticum, and M. magneticum, and 6.3, 3.3, and 2.0 mg magnetite per liter per day were obtained from each species, respectively (Heyen and Schu¨ler 2003). Using a similar type of fermenter, except that dissolved oxygen was controlled to an optimal level using the change of cell growth rate rather than from a direct measurement from the sensitive oxygen electrode, Sun et al. (2008) obtained a cell density of OD565 of 7.24 for M. gryphiswaldense after 60 h of culture. The cell yield (dry weight) was 2.17 g/l, and the yield of magnetosomes (dry weight) was 41.7 mg/l and 16.7 mg/l/day. By decreasing the amount of carbon and electron source (lactate) in the same fermenter, Liu et al. (2010) reported later growth and magnetosome yields of OD565nm of 12 and 55.49 mg/l/day, respectively, after 36 h of culture again using M. gryphiswaldense.

Applications of Cells of Magnetotactic Bacteria Both live and dead magnetotactic bacterial cells have proven useful in medical, magnetic, and environmental applications. They have been used to magnetically separate granulocytes and monocytes after having been phagocytized by them (Matsunaga et al. 1989). Because of the relatively easy separation of magnetic cells, the use of magnetotactic bacteria in the uptake and remediation of heavy metals and radionucleotides from wastewater has been discussed and investigated (Bahaj et al. 1993, 1998a, b, c; Arakaki et al. 2002). Cells of polar magnetotactic bacteria have been used to determine south magnetic poles in meteorites and rocks containing fine-grained ( Fig. 12.23) (Chang and Kirschvink 1989; Chang et al. 1989) and in the martian meteorite ALH84001 (Thomas-Keprta et al. 2000, 2001, 2002; Clemett et al. 2002). These crystals, referred to as ‘‘magnetofossils,’’ have been used as evidence for the past presence of magnetotactic bacteria in sediments and in meteorite ALH84001. The presence and interpretation of these crystals in martian meteorite ALH84001, in particular, have instigated great controversy and debate. If the magnetite crystals were indeed biogenic, the implication was that bacterial life had existed on ancient Mars (McKay et al. 1996; Thomas-Keprta et al. 2000, 2001, 2002; Buseck et al. 2001; Clemett et al. 2002; Weiss et al. 2004). In turn, this debate has led to a number of criteria to be used to distinguish biogenic magnetite from inorganically produced magnetite (Thomas-Keprta et al. 2000; Arato´ et al. 2005;

We are grateful to the continued collaboration, support, and encouragement of R. B. Frankel. DAB is supported by US National Science Foundation (NSF) Grant EAR-0920718. CTL is supported by a grant from the Fondation pour la Recherche Me´dicale SPF20101220993. DS has been supported by grants from the Deutsche Forschungsgemeinschaft, the German Bundesministerium fu¨r Bildung und Forschung, and the European Union.

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13 Luminous Bacteria Paul V. Dunlap1 . Henryk Urbanczyk2 1 University of Michigan, Ann Arbor, MI, USA 2 University of Miyazaki, Miyazaki City, Miyazaki, Japan

Introduction and Historical Perspective . . . . . . . . . . . . . . . . . . 495 Biochemistry of Bacterial Luminescence . . . . . . . . . . . . . . . . . . 496 Species and Phylogeny of Luminous Bacteria . . . . . . . . . . . . . 498 The Bacterial Luminescence Operon . . . . . . . . . . . . . . . . . . . . . . 500 Genomes of Luminous Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . 502 Evolutionary Origin and Function of Bacterial Luminescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 502 Horizontal Acquisition of the Bacterial lux Genes . . . . . . . . 504 Habitats and Ecology of Luminous Bacteria . . . . . . . . . . . . . . 505 Marine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 505 Freshwater . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 506 Terrestrial . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 507 Parasitism of Marine Invertebrates . . . . . . . . . . . . . . . . . . . . . . 507 Parasitism of Vertebrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 508 Bioluminescent Symbiosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 508 Patterns of Host Affiliation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 511 Species Specificity, Cosymbiosis, and Symbiont: Host Codivergence . . . . . . . . . . . . . . . . . . . . . . . . . . . 511 Symbiont Acquisition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 513 Quorum Sensing Control (Autoinduction) of Bacterial Luminescence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 514 Isolation, Cultivation, Storage, and Identification of Luminous Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 518 Isolation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 518 Cultivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 519 Storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 519 Identification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 519 Abstract Luminous bacteria are those bacteria that carry the lux genes, genes that code for proteins involved in light production. Many luminous bacteria emit light at high, easily visible levels in laboratory culture and in nature, and the phenomenon of light emission has generated interest in these bacteria for over 125 years. Luminous bacteria are especially common in ocean environments where they colonize a variety of habitats, but some

species are found in brackish, freshwater, and terrestrial environments. This chapter, which begins with an historical perspective, summarizes current understanding of the biochemistry and genetics of bacterial light emission, the taxonomy and phylogenetics of light-emitting bacteria, the evolutionary origins and hypothesized physiological and ecological functions of bacterial luminescence, the distributions and activities of these bacteria in nature, their symbiotic interactions with animals and especially with marine fishes, and the quorum sensing regulatory circuitry controlling light production at the operon level. This chapter concludes with information on the isolation, cultivation, storage, and identification of luminous bacteria.

Introduction and Historical Perspective Luminous bacteria are those bacteria that carry lux genes, at a minimum luxA and luxB, the genes coding for bacterial luciferase, either as vertically inherited genes or genes naturally acquired by horizontal transfer. Most of the currently known luminous bacteria express the lux genes and produce light at high, readily visible levels in laboratory culture (> Fig. 13.1) or in nature. Not all lux gene-carrying bacteria, however, produce levels of light visible to the human eye. To date, luminous bacteria have been found in only three closely related Gammaproteobacteria families, Vibrionaceae, Enterobacteriaceae, and Shewanellaceae, and most species are members of Vibrionaceae. Most luminous bacteria are facultatively aerobic, but two, Shewanella hanedai (Jensen et al. 1980) and Shewanella woodyi (Makemson et al. 1997), are respiratory. Additional and detailed information on the metabolism, physiology, morphology, and ecology of these bacterial groups and individual species can be found in Baumann and Baumann (1981), Baumann et al. (1984), Farmer and Hickman–Brenner (1992), Boemare et al. (1993), Forst et al. (1997), and Urbanczyk et al. (2007). Bacterial luminescence is one of several evolutionarily distinct forms of bioluminescence, an attribute of a wide diversity of eukaryotic organisms (Hastings 1995; Widder 2010). The ability of certain bacteria to produce light has been known since 1875, when Pflu¨ger (1875) related the luminescence coming from the slime of fish to bacteria present in the slime (Harvey 1957; Robertson et al. 2011). Many earlier observations suggest the presence of luminous bacteria and knowledge of their existence. 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 emit light (Harvey 1940, 1952). Many

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. Fig.13.1 Bacterial bioluminescence. Colonies of P. mandapamensis from the light organ of the cardinalfish Siphamia versicolor (Perciformes: Apogonidae) are shown growing on LSW-70 agar plates. The plate was photographed in the dark by the light produced by the bacteria

years before those observations and long before bacteria and the oxygen dependence of bacterial luminescence were known, 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 or in marine and terrestrial animals. According to Harvey (1940), J. F. Heller in 1854 was the first to give a name, Sarcina noctiluca, to an organism suspected to be responsible for luminescence. Following Pflu¨ger’s work in 1875, other scientists working in the late 1800s and early 1900s isolated and named luminous bacteria, including ‘‘Bacterium lucens,’’ ‘‘Micrococcus phosphorescens,’’ ‘‘Micrococcus pflu¨geri,’’ ‘‘Bacillus phosphorescens,’’ and ‘‘Bacterium phosphoreum’’ (Neush 1879; Ludwig 1884; Fischer 1887; Molisch 1912; Dahlgren 1915; Zobell 1946; Harvey 1952, 1957; Robertson et al. 2011). Particularly notable among early researchers of bacterial luminescence was Martinus W. Beijerinck, a founder of general microbiology, who carried out research on the physiology of light-emitting bacteria and who coined the name Photobacterium, a genus within which he grouped all luminous bacteria (Beijerinck 1889a, b, 1891, 1916; van Iterson et al. 1940; Robertson 2003; Robertson et al. 2011). The recent revival and phylogenetic characterization of strains isolated by Beijerinck and stored in the 1920s (Figge et al. 2011) provide a direct link to the origins of general microbiology and the first studies of luminous bacteria.

Following these early studies at the end of the nineteenth and the beginning of the twentieth century, luminous bacteria were isolated from various habitats, the chemistry of bacterial light production and the culture requirements for growth and luminescence of the bacteria were characterized, and they were placed taxonomically as microbial systematics developed (e.g., Zobell and Upham 1944; Farghaly 1950; Johnson 1951). In the latter half of the twentieth century and continuing to date, taxonomic efforts have paralleled the growth of microbiology, incorporating the tools and knowledge developing from advances in biochemistry, physiology, and genetics (Baumann and Baumann 1977, 1981; Farmer and Hickman–Brenner 1992; Hastings and Nealson 1977, 1981; Hendrie et al. 1970; Nealson and Hastings 1992; Singleton and Skerman 1973). Presently, over 25 species of luminous bacteria are validly described (> Table 13.1). Marine luminous species are found in Aliivibrio, Photobacterium, Vibrio (Vibrionaceae), and Shewanella (Shewanellaceae), and terrestrial light-producing species are members of Photorhabdus (Enterobacteriaceae) (Dunlap and Kita–Tuskamoto 2006; Urbanczyk et al. 2007, 2008; Ast et al. 2009; Dunlap 2009; Yoshizawa et al. 2009a, b; 2010a, b). Current understanding of the systematic relationships of luminous bacteria, as well as recent descriptions of new species, has utilized phylogenetic analysis of multiple, functionally independent housekeeping genes, including the 16S rRNA gene, gyrB, pyrH, and recA, among others (e.g., Ast and Dunlap 2005; Thompson et al. 2005; Ast et al. 2007b, 2009; Urbanczyk et al. 2007). Particularly useful for resolving the separate species status of closely related luminous bacteria is sequence analysis of the lux genes, luxCDABE, found to date in all luminous bacteria, due to their relatively rapid sequence divergence compared to most housekeeping genes (Ast and Dunlap 2004, 2005; Dunlap et al. 2004; Ast et al. 2007a; Urbanczyk et al. 2007). The description of several new species in the past few years (> Table 13.1) suggests that many more species of luminous bacteria remain to be discovered. The advents of whole genome sequencing, metagenomics, and single-cell genomics and their application to luminous bacteria will undoubtedly provide additional insight into the systematics of luminous bacteria, the evolution of the bacterial luminescence system, and many other aspects of the biology of these bacteria.

Biochemistry of Bacterial Luminescence 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 (fatty) aldehyde (RCHO) by O2 to produce blue-green light according to the following reaction: FMNH2 þ O2 þ RCHO

luciferase

! FMN þ H2 O þ RCOOH þ hn ð490 nmÞ

In the luminescence reaction, binding of FMNH2 by the enzyme is followed by interaction with O2 to form a flavin-4ahydroperoxide. Association of this complex with aldehyde forms

Luminous Bacteria

13

. Table 13.1 Species and habitats of luminous bacteria Species

Habitatsa

Selected references

Marine Aliivibrio fischeri

Coastal seawater,

Boettcher and Ruby (1990), Fitzgerald (1977), Lee and Ruby (1992)

Light organs of squid and fish

Reichelt and Baumman (1973), Ruby and Nealson (1976)

logei

Coastal seawater, sediment

Ast et al. (2009), Bang et al. (1978), Urbanczyk et al. (2007)

salmonicida

Tissue lesions of Atlantic salmon

Hjerde et al. (2008), Nelson et al. (2007), Urbanczyk et al. (2007)

sifiae

Coastal seawater

Ast et al. (2009), Yoshizawa et al. (2010a)

‘‘thorii’’

Light organs of squid

Ast et al. (2009), Fidopiastis et al. (1998)

wodanis

Coastal seawater, diseased farmed salmon, light organs of squid

Ast et al. (2009), Lunder et al. (2000), Urbanczyk et al. (2007)

Coastal seawater

Yoshizawa et al. (2009b)

damselae

Coastal seawater

Smith et al. (1991), Urbanczyk et al. (2008)

kishitanii

Light organs and skin of fish

Ast and Dunlap (2005), Ast et al. (2007)

leiognathi

Coastal seawater, light organs of fish Boisvert et al. (1967), Dunlap et al. (2007), Dunlap et al. (2008), Fukasawa and Dunlap (1986), Fukasawa et al. (1988), Reichelt et al. (1977)

mandapamensis

Coastal seawater, light organs of fish Hendrie et al. (1970), Kaeding et al. (2007), Reichelt and Baumman (1973), Urbanczyk et al. (2011b), Wada et al. (2006)

phosphoreum

Coastal and pelagic seawater

Ast and Dunlap (2005), Ast et al. (2007a), Baumann et al. (1980), Budsberg et al. (2003), Ruby and Morin (1978), Wimpee et al. (1991)

Photodesmus katoptronb

Light organs of anomalopid fish

Haygood (1990), Hendry and Dunlap (2011), Wolfe and Haygood (1991)

Shewanella hanedai

Seawater and sediment

Jensen et al. (1980)

woodyi

Seawater and squid ink

Makemson et al. (1997)

Coastal seawater

Yoshizawa et al. (2009a)

Ruby and Nealson (1978), Ruby et al. (2005), Urbanczyk et al. (2007)

Photobacterium aquimaris

Candidatus

Vibrio azureus ‘‘beijerinckii’’

Coastal seawater

Figge et al. (2011)

campbellii

Coastal seawater

Lin et al. (2010)

chagasii

Coastal seawater, surfaces and intestines of marine animals

Thompson et al. (2003), Urbanczyk et al. (2008)

harveyi

Coastal seawater, sediment

Gomez–Gil et al. (2004), O’Brien and Sizemore (1979), Reichelt and Baumman (1973), Ruby and Nealson (1978), Yetinson and Shilo (1979)

mediterraneac

Coastal seawater

Pujalte and Garay (1986), Ortiz Conde et al. (1989)

orientalis

Seawater, surface of shrimp

Yang et al. (1983)

sagamiensis

Coastal seawater

Yoshizawa et al. (2010b)

splendidus

Coastal seawater

Baumann et al. (1980), Nealson et al. (1993)

vulnificus

Coastal seawater, oysters

Oliver et al. (1986), Urbanczyk et al. (2008)

Estuaries, bays coastal seawater

Kaeding et al. 2007; Palmer and Colwell (1991), Ramaiah et al., (2000), Zo et al. (2009)

Human skin lesions

Farmer et al. (1989); Fisher–Le Saux et al. (1999), Peel et al. (1999)

Brackish/Estuarine Vibrio cholerae Terrestrial Photorhabdus asymbiotica

497

498

13

Luminous Bacteria

. Table 13.1 (continued) Habitatsa

Selected references

luminescens

Insect larvae infected with heterorhabditid nematodes

Boemare et al. (1993), Fisher–Le Saux et al. (1999)

temperata

Insect larvae infected with heterorhabditid nematodes

Fisher–Le Saux et al. 1999

Species

a

Representative habitats of luminous strains are listed. Candidatus name, not cultured (Hendry and Dunlap 2011). c Ability of this species to luminescence is not well established; the single strain reported as luminous (Pujalte and Garay 1986) may not be available. b

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 0.1–1.0 photons. The reaction is highly specific for FMNH2, and the aldehyde substrate in vivo is likely to be tetradecanal. FMNH2 is provided by the activity of an NAD(P)H-flavin oxidoreductase (flavin reductase). Synthesis of the long-chain aldehyde is catalyzed by a fatty acid reductase complex composed of three polypeptides, an NADPH-dependent acyl protein reductase (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. Luciferases from different species of luminous bacteria exhibit substantial amino acid residue and nucleotide sequence identity (Meighen and Dunlap 1993; Dunlap et al. 2007), consistent with a common evolutionary origin of luminescence in bacteria. For references and detailed information on the biochemistry of bacterial light production, the reader is directed to reviews by Hastings (1995), Lee et al. (1990), Hastings et al. (1985), Meighen (1988; 1991), Meighen and Dunlap (1993), and Wilson and Hastings (1998).

Species and Phylogeny of Luminous Bacteria Over 25 species of luminous bacteria are validly described at this time (> Table 13.1). Taxonomically, luminous bacteria are members of six of genera in three Gammaproteobacteria families: Vibrionaceae, Enterobacteriaceae, and Shewanellaceae (> Fig. 13.2) To date, no luminous strains belonging to other families have been reported. Most luminous species are members of Aliivibrio, Vibrio, and Photobacterium in Vibrionaceae. Detailed phylogenetic analysis has shown that most extant luminous members of Vibrionaceae acquired their luxCDABE genes vertically, with only a few cases of acquisition by intraspecies horizontal transfer from members of Vibrionaceae, whereas luminous members of Enterobacteriaceae and Shewanellaceae apparently acquired their lux genes by horizontal transfer from members of Vibrionaceae (Urbanczyk et al. 2008). These considerations, and others described below, suggest that the luxCDABE-based luminescence system of bacteria arose just once evolutionarily, apparently in an ancestor of Vibrionaceae (Urbanczyk et al. 2008). Several new species of luminous bacteria have been described in the past few years (> Table 13.1, > Fig. 13.2).

These include Aliivibrio sifiae (Ast et al. 2009; Yoshizawa et al. 2010a), Photobacterium kishitanii (Ast et al. 2007a), Photobacterium aquimaris (Yoshizawa et al. 2009b), Candidatus Photodesmus katoptron (Hendry and Dunlap 2011), Vibrio azureus (Yoshizawa et al. 2009a), and Vibrio sagamiensis (Yoshizawa et al. 2010b). Recent studies have also revealed the presence of luminous strains of species not previously reported as luminous, that is, Vibrio campbellii (Lin et al. 2010), Vibrio vulnificus (Urbanczyk et al. 2008), and Vibrio damsela (Urbanczyk et al. 2008). With respect to V. campbellii, a recent genomic analysis has revealed strain ATCC BAA-1116 (aka BB120), previously classified as Vibrio harveyi and studied intensively for quorum sensing control of luminescence and other cellular functions in this species (e.g., Bassler et al. 1993; Waters and Bassler 2005; Long et al. 2009), is actually a member of V. campbellii (Lin et al. 2010). Furthermore, newly recognized clades, for example, Aliivibrio ‘‘thorii’’ and Vibrio ‘‘beijerinckii’’ (Ast et al. 2009; Figge et al. 2011), have been identified, and formal description of these and other new species is under way. In most cases, taxonomic identification has followed cultivation-based detection of light emission; most of the bacteria listed in > Table 13.1 grow and emit light in laboratory media. However, several luminous bacteria, bioluminescent symbionts of anomalopids (flashlight fish) and ceratioids (deep-sea anglerfish), are known that have not been cultured; these bacteria are members of Vibrionaceae but are divergent from known species of luminous bacteria (Haygood 1990, 1993; Haygood and Distel 1993). Very recently, luminous bacteria symbiotic with the anomalopid fish Anomalops katoptron were characterized phylogenetically and assigned Candidatus status as a new Vibrionaceae genus and species, Photodesmus katoptron (Hendry and Dunlap 2011). Most luminous strains isolated from natural habitats group taxonomically as members of well-recognized species that typically are considered to be luminous (> Table 13.1). However, luminescence often is not a uniformly consistent phenotype of even these luminous species (e.g., Wollenberg et al. 2011) or their genera. Nonluminous species of Vibrio are well known and are more common than luminous strains (e.g., Baumann and Baumann 1981), and several nonluminous species of Photobacterium and Aliivibrio have been found, some of which apparently lack lux genes (Dunlap and Ast 2005; Urbanczyk et al. 2011a). Furthermore, strains of Photorhabdus luminescens symbiotic with entomopathogenic nematodes have been found that do not produce light and lack genes necessary for light

Luminous Bacteria

13

Enterobacteriaceae

Xenorhabdus nematophila Photorhabdus asymbiotica Ph. luminescens Ph. temperata Escherichia coli Shewanella oneidensis Shewanellaceae S. cloweliana S. hanedai S. woodyi Vibrionaceae Vibrio mimicus V. cholerae V. diazotrophicus V. vulnificus V. splendidus V. cyclitrophicus V. lentus V. chagasii V. mediterranei V. orientalis V. pelagius V. alginolyticus V. parahaemolyticus V. azureus

V. sagamiensis V. harveyi V. campbelli

80 changes

Aliivibrio fischeri A. finisterrensis A. logei A. salmonicida A. sifiae A. wodanis A. “thorii” Photobacterium damselae P. profundum P. leigonathi P. mandapamensis P. angustum P. aquimaris P. kishitanii P. phospohoreum P. iliopiscarium

. Fig. 13.2 Phylogeny of luminous bacteria. The analysis, parsimony implemented in PAUP*, is based on sequences of the 16S rRNA and gyrB genes. Luminous species (in boldface) are found in three families, Vibrionaceae, Shewanellaceae, and Enterobacteriaceae. These families contain many more nonluminous species than shown here. Also, recently identified luminous bacteria, for example, Vibrio ‘‘beijerinckii’’ (proposed name) (Figge et al. 2011) and Candidatus Photodesmus katoptron (Hendry and Dunlap 2011), and additional species whose descriptions are underway, are not shown

production (Akhurst and Boemare 1986; Forst and Nealson 1996). In addition, strains luminous on primary isolation often become dim or dark in laboratory culture (Nealson and Hastings 1979, 1992; Akhurst 1980; Silverman et al. 1989), and some species that grow well in laboratory culture at room temperature, that is, A. logei and S. hanedai, typically produce readily visible light only when grown at cooler temperatures. In some cases, that is, luminous bacteria infecting crustaceans (Giard and Billet 1889b) and strains of A. fischeri symbiotic with the Hawaiian sepiolid squid, Euprymna scolopes (Boettcher and Ruby 1990), the bacteria produce a high level of light in their

natural habitat but produce little or no light when grown in laboratory culture. Adding to this complexity, V. cholerae, generally considered to be a nonluminous species, has many luminous strains (e.g., Kaeding et al. 2007; Zo et al. 2009), and many of the nonluminous strains of this species carry lux genes that apparently are not expressed in laboratory culture (Palmer and Colwell 1991; Ramaiah et al. 2000). In addition, bacteria identified as related to V. harveyi and V. cincinnatiensis carry the lux genes but have been found to have lux gene mutations that result in a dark phenotype (O’Grady and Wimpee 2008). Furthermore,

499

500

13

Luminous Bacteria

luminous strains of three other species generally known as nonluminous, Vibrio vulnificus, Vibrio chagasii, and Photobacterium damselae, recently were identified (Urbanczyk et al. 2008). This substantial variation in the incidence of luminous strains within a species has implications for understanding the evolutionary origins of bacterial luminescence and its patterns of inheritance, as described in sections that follow. It should be noted that the presence of luminescence strains has likely been overlooked in many species. Routine use of cooler temperatures (10–20  C) for growth and examination, and utilization of conditioned media, inducers, and luciferase substrates (Fidopiastis et al. 1999), along with the application of probes for luxA and other lux genes (Wimpee et al. 1991), and full sequence-based characterization of the lux operons of new luminous bacteria will undoubtedly lead to a more complete understanding of the species diversity of bacteria able to produce light. A further complication in gaining a more comprehensive understanding of the diversity of luminous bacteria is that descriptions of new species of luminous bacteria often lack detailed information on the lux genes, relationships to type strains, or detailed phylogenetic analysis (e.g., Yoshizawa et al. 2009b, 2010b). In this regard, characterization of multiple newly isolated strains, the use of multiple independent loci, and the use of type strains and other key strains are imperatives in new species descriptions for revealing the otherwise hidden species diversity of luminous bacteria (e.g., Ast et al. 2009).

The Bacterial Luminescence Operon The genes coding for the a- and b-subunits of bacterial luciferase, luxA and luxB, respectively, are part of the lux operon, luxCDABE, which is present in the genomes of all luminous bacteria examined to date as a conserved, contiguous, and coordinately expressed set of genes (> Fig. 13.3). The luxC, luxD, and luxE genes, respectively, code for the r, s, and t polypeptides of the fatty acid reductase complex that synthesizes and recycles aldehyde substrate for luciferase. The lux operons of most bacteria also contain luxG, which codes for a flavin reductase (Lin et al. 1998; Meighen and Dunlap 1993; Nijvipakul et al. 2008; Swartzman et al. 1990a). The absence of luxG from the lux operon of Ph. luminescens apparently is compensated for by the activity of a flavin reductase activity coded for by an Escherichia coli fre-like gene, homologs of which are found in various luminous bacteria (Zenno et al. 1992, 1994; Zenno and Saigo 1994). Several species of Photobacterium bear an additional lux operon gene, luxF, between luxB and luxE. The luxF gene, coding for a nonfluorescent flavoprotein, is apparently specific to Photobacterium, as it is not present in the lux operons of Aliivibrio, Photorhabdus, Shewanella, or Vibrio species (> Fig. 13.3), but it has been secondarily lost in Photobacterium leiognathi (Ast and Dunlap 2004). The LuxF protein might function in the luminescence system by scavenging an inhibitory side product of the luciferase reaction (Moore and James 1995), but it is not necessary for light production even in those Photobacterium species that normally carry this gene (Kaeding

et al. 2007). In the examined species of luminous Photobacterium, the lux operon genes are followed, without a transcriptional stop or other regulatory sites, by genes involved in the synthesis of riboflavin, ribEBHA, the products of which presumably function in generating FMNH2, a substrate of luciferase (Lee and Meighen 1992; Lee et al. 1994; Lin et al. 2001; Sung and Lee 2004; Ast et al. 2007b). We refer to this gene arrangement as the Photobacterium lux-rib operon (> Fig. 13.3). Strains of P. phosphoreum lack one of the rib genes, ribE; the gene presumably was lost in the divergence from an ancestral Photobacterium that gave rise to this species. The presence of genes for synthesis of riboflavin as part of the lux operon might facilitate light production by ensuring coordinate synthesis of luciferase and substrates for the enzyme. In this regard, the lux operon of V. campbellii (previously classified as V. harveyi; Lin et al. 2010) contains ribB, coding for 3,4-dihydroxy-2-butanone 4-phosphate synthase, a key enzyme in riboflavin synthesis (referred to originally as luxH; Swartzman et al. 1990b) as the final gene, as does the lux operon of Candidatus Photodesmus katoptron (Hendry and Dunlap 2011). Furthermore, although ribB is not part of the lux operon of A. fischeri, its expression nonetheless is under the same quorum sensing control as the lux genes (Callahan and Dunlap 2000). In addition to presence of ribEBHA genes in Photobacterium as part of the lux-rib operon, genes upstream of the lux operon contribute to luminescence and also show genus and species differences. In Photobacterium mandapamensis, for example, the lux-rib operon is preceded by lumQ and lumP, which form the lumazine operon. The function of lumQ is not yet known, although it might code for a DNA binding protein (Lin et al. 1995). LumP, a 21 kDa fluorescent accessory protein referred to as lumazine protein, functions to shift the emission wavelength of luciferase from blue-green (495 nm) to blue (475–486 nm) and enhance the intensity of light emission (Lee 1993; O’Kane et al. 1985,1991; Petushkov et al. 1996). LumP, which has been isolated from P. phosphoreum and strains called P. leiognathi (which actually are P. mandapamensis, see below) and also purified from P. kishitanii, contains a noncovalently bound fluorophore, 6,7-dimethyl-8-ribityllumazine, the immediate biosynthetic precursor of riboflavin (O’Kane et al. 1985; Sato et al. 2010; Small et al. 1980). In P. leiognathi lumP is not found, although approximately 200 nucleotides of the P. leiognathi luxC–lumQ intergenic region can be aligned to the P. mandapamensis lumP gene (> Fig. 13.4; Ast et al. 2007b). The activity of the LumP protein apparently accounts for the blueshifted luminescence of P. mandapamensis compared to P. leiognathi, one of the diagnostic traits distinguishing these two species (O’Kane et al. 1985, 1991; Lee 1993; Petushkov et al. 1996; Ast and Dunlap 2004; Kaeding et al. 2007). The genes flanking the P. leiognathi and P. mandapamensis lux-rib operons are homologous to a single contiguous region in nonluminous P. angustum (Lin et al. 1993, 1995, 1996a, b, 2001; Ast et al. 2007b). In the examined Aliivibrio species, regulatory genes, luxI and luxR, which control transcription of the lux operon, precede or flank the luxCDABEG genes (> Fig. 13.3). The luxI gene codes for an acyl-homoserine lactone (acyl-HSL) synthase

Luminous Bacteria luxR luxI

luxC

luxD

luxA

luxB

luxE

luxG

13

Aliivibrio fischeri luxR1

luxC

luxD

luxA

luxB

luxE

luxG luxR2 luxI

luxR1

luxC

luxD

luxA

luxB

luxE

luxG luxR2 luxI

luxI

luxC

luxD

luxA

luxB

luxE

luxG

luxC

luxD

luxA

luxB

luxE

luxG

luxC

luxD

luxA

luxB

luxC

luxD

luxA

luxB

luxC

luxD

luxA

luxB

luxF

luxE

luxC

luxD

luxA

luxB

luxF

luxE

luxC

luxD

luxA

luxB

luxE

luxG

luxC

luxD

luxA

luxB

luxE

luxG

luxC

luxD

luxA

luxB

luxE

luxG

luxC

luxD

luxA

luxB

luxE

Aliivibrio logei

Aliivibrio salmonicida luxR

Shewanella hanedai

Shewanella woodyi

luxF

luxE

luxG

ribE

ribB ribH

ribA

Photobacterium kishitanii luxE

luxG

ribE ribB

ribH

ribA

luxG

ribE

ribB

ribH

luxG

ribB ribH

Photobacterium leiognathi ribA

Photobacterium mandapamensis ribA

Photobacterium phosphoreum

ribB

Vibrio campbellii

Vibrio cholerae ribB

Photodesmus katoptron

Photorhabdus luminescens

. Fig. 13.3 Genes of the lux operons of luminous bacteria. Shown are the lux genes and the organization of lux operons for those bacteria for which complete lux operon sequence data are available. Contiguous genes of the luminescence operons of luminous bacteria are aligned to highlight commonalities and differences. Four distinct types of lux operons are evident based on commonalities of gene content, organization, and sequence similarity, (1) the Aliivibrio/Shewanella type, with luxI/luxR regulatory genes; (2) Photobacterium type, with ribEBHA genes; (3) the Vibrio/Candidatus Photodesmus type, with neither regulatory nor additional linked genes; and (4) the Photorhabdus type, composed of just the core luxCDABE genes

(Schaefer et al. 1996), and luxR codes for a receptor protein that interacts with acyl-HSL to activate transcription of the lux operon (Engebrecht et al. 1983), as described in more detail below. In Aliivibrio fischeri, luxI is the first gene of the lux operon, and luxR, upstream of luxI, is divergently transcribed (> Fig. 13.3). The same gene arrangement is present in Shewanella hanedai, and this identity together with the high degree of lux gene sequence similarity in S. hanedai and A. fischeri has led to the suggestion that S. hanedai acquired its lux operon by horizontal transfer from A. fischeri or the ancestor of A. fischeri (Urbanczyk et al. 2008), as described in more detail

below. In Aliivibrio salmonicida, a bacterium that requires exogenous addition of aldehyde to produce a high level of light (Fidopiastis et al. 1999), two luxR genes, homologous to A. fischeri luxR, flank the lux operon; a luxI gene also is present, divergently transcribed from the downstream luxR (> Fig. 13.3; Nelson et al. 2007; Hjerde et al. 2008). Very recently, the same arrangement of luxI and luxR genes as in A. salmonicida was identified in Aliivibrio logei (Manukhov et al. 2011). In contrast to A. salmonicida, however, A. logei does not require exogenous aldehyde to produce a high level of light (Manukhov et al. 2011); sequence comparison of the two operons identified mutations in

501

502

13

Luminous Bacteria

a

putA

lumQ

b

putA

lumQ

c

transposase

lumP

luxC

luxC

lumQ

luxC

1kb

. Fig. 13.4 Region upstream of the lux operon in Photobacterium. (a) The luxrib operon is preceded in P. mandapamensis by lumQ and lumP. (b) For the lux-rib1 operon of P. leiognathi, lumQ is present upstream and lumP is not found, although approximately 200 nucleotides of the P. leiognathi luxC–lumQ intergenic region can be aligned to the P. mandapamensis lumP gene sequence. (c) The region upstream of the lux-rib2 operon of P. leiognathi contains lumQ and a transposase gene. For details, see the text and Ast et al. (2007b). The regions flanking the lux-rib operons of other Photobacterium species remain to be defined, but preliminary information for some species indicates differences from the arrangement shown here (Urbanczyk et al. unpublished data)

luxD of A. salmonicida that presumably account for the exogenous aldehyde requirement of this species. Genes flanking the lux operons of other luminous Aliivibrio species (> Table 13.1) apparently have not yet been identified. The arrangement of genes flanking the lux operons of the examined Vibrio species differs substantially from that in Photobacterium and in Aliivibrio (> Fig. 13.3). First, regulatory genes controlling transcription of the lux operon are not part of and are not adjacent to the lux operon in those Vibrio species examined; specifically, a luxR gene, which is not homologous to A. fischeri luxR, is not physically associated with the lux operon in V. campbellii (V. harveyi); the role of luxRVh in the phosphorelay cascade controlling luminescence in this species is outlined below. Conservation of luxCDABE as a unit might reflect a need for close interaction of luciferase and fatty acid reductase proteins, based on coordinate regulation, to facilitate substrate generation necessary for efficient light production. However, it is not obvious what led to the genus-specific differences in the presence of genes flanking and contiguous with the lux operons of luminous members of Aliivibrio, Photobacterium, and Vibrio, three closely related genera of Vibrionaceae.

Genomes of Luminous Bacteria The genomes of luminous Vibrio and Photobacterium species are similar in structure, overall size, and organization to other members of Vibrionaceae, with two chromosomes of unequal size and an overall size of approximately 4.5–5.4 Mb (Egan et al. 2005; Okada et al. 2005; Ruby et al. 2005; Vezzi et al. 2005; Reen et al. 2006; Lauro et al. 2009; Ast et al. 2007a;

Urbanczyk et al. 2011a, b; Urbanczyk et al. unpublished data). The significance of this organization for members of Vibrionaceae is not yet known, but differences between the two replicons suggest that each chromosome carries out substantially different roles in the cell. More of the core (essential) genes are found on the large chromosome, whereas the small chromosome contains mostly lineage specific genes. Furthermore, gene content and position appear to be more highly conserved on the large chromosome than on the small chromosome (Reen et al. 2006). The small chromosome in members of Vibrionaceae nonetheless contains some essential genes (Reen et al. 2006), which might guarantee retention of the small chromosome during cell division (Egan et al. 2005). The origin of the small chromosome in Vibrionaceae remains unknown but has been hypothesized in V. cholerae to have originated from a plasmid that accumulated additional genes, including some genes transferred from the large chromosome (Egan et al. 2005). The apparent ubiquity of two chromosomes of unequal size in Vibrionaceae suggests that the small chromosome may have arisen in the ancestral lineage leading to Vibrionaceae. In view of the different predicted roles for the large and small chromosomes in Vibrionaceae, it is significant that in luminous species for which complete sequences of both chromosomes are available, the lux operons are all located on the small chromosome. These species include A. fischeri (Ruby et al. 2005; Mandel et al. 2009), A. salmonicida (Hjerde et al. 2008), and V. campbellii (Lin et al. 2010). This pattern indicates that the lux genes have an accessory function, that is, they are not part of the core genome, a view that is consistent with the many nonluminous Vibrionaceae species and many nonluminous strains of luminous species, as noted above. Which chromosome, large or small, carries the lux genes is not yet known in Photobacterium species. A single chromosome is characteristic of Enterobacteriaceae and Shewanellaceae, and the lux genes in Photorhabdus and luminous Shewanella are chromosomal. The genomes of luminous bacteria analyzed to date have been found to carry multiple rRNA operons. Specifically, the genome of A. fischeri carries 12 rRNA operons (Ruby et al. 2005), the P. kishitanii genome has eight or more (Ast et al. 2007a), the P. mandapamensis genome has six or more (Urbanczyk et al. 2011b), and the Ph. luminescens genome has seven (Duchaud et al. 2003; Wilkinson et al. 2009). A high copy number of rRNA operons may be an adaptation for a copiotrophic lifestyle and for rapid response to nutrient availability (Klappenback et al. 2000; Lauro et al. 2009). Taken together, the relatively large genome size and multiple rRNA operons of luminous bacteria and other members of Vibrionaceae may be adaptations for rapidly utilizing a wide range of different nutrients under feast-or-famine conditions.

Evolutionary Origin and Function of Bacterial Luminescence The origin of bacterial luminescence has been of interest since the early days of microbiology. The natural presence of genes

Luminous Bacteria

necessary for producing light defines the luminous bacteria. The necessary genes, luxA and luxB, encoding the luciferase subunits, luxC, luxD and luxE, for the fatty acid reductase subunits, and luxG, encoding a flavin reductase, are consistently found together as a cotranscribed unit, luxCDABEG. The reason for this conservation of lux genes as a unit is not known, but it might relate to efficient light production; the contiguous presence of these genes as an operon might help promote the coordinated production of luciferase and substrates for luciferase, long-chain aldehyde and reduced flavin mononucleotide (FMNH2). The conservation of these genes as a unit in nearly all luminous bacteria examined suggests that the lux operon arose just once in the distant past. Supporting this view, phylogenetic analysis demonstrates that the individual lux genes of different bacterial species are homologous, as was suggested by the high levels of amino acid sequence identities of the inferred Lux proteins. This homology implies that the bacterial luxCDABEG genes arose one time in the evolutionary past. The use by luciferase of oxygen as a substrate implies that this enzymatic activity originated after oxygenic photosynthesis by ancestors of modern-day cyanobacteria began to increase the level of O2 on Earth, approximately 2.4 billion years ago, during the Great Oxidation Event. A marine origin for bacterial luminescence (Palmer and Colwell 1991; Dunlap 2009) seems likely because most species of luminous bacteria are marine (> Table 13.1). Seliger (1987) proposed that bacterial luminescence arose under ecological selection for light emission. 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 flavin-dependent, aldehyde-oxidizing protoluciferase produced sufficient light and with an appropriate emission spectrum, to be detected by phototactic organisms. Ingestion by visually cueing animals of particles colonized and made luminous by these early luminous bacteria presumably enhanced their reproduction by bringing them into the animal’s nutrient-rich digestive system, ensuring the emitter’s survival and thereby possibly leading to selection for more intense light output (Widder 2010). 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). An alternative hypothesis for the evolution of bacterial luciferase for DNA repair (Czyz˙ et al. 2003) has been called into question (Walker et al. 2006). A single gene was hypothesized to encode bacterial protoluciferase (O’Kane and Prasher 1992). Although a singlesubunit protoluciferase, monomer or dimer, presumably would have differed somewhat from the modern-day luciferase a-subunit and therefore might have produced light, the inability of either of the extant 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 protoluminescence may have arisen following a gene duplication event that is postulated to

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have created luxB from luxA (Baldwin et al. 1979; O’Kane and Prasher 1992; Meighen and Dunlap 1993). Based on amino acid sequence identities, a tandem duplication of the ancestral luxA gene, followed by sequence divergence in the duplicated gene, is thought to have given rise to luxB, leading to the formation of the heterodimeric luciferase present in extant luminous bacteria. Similarly, a tandem duplication of luxB followed by loss of approximately 300 nucleotides coding for N-terminus amino acids is thought to have given rise to luxF in a luminescent ancestor of Photobacterium; this gene apparently was later secondarily lost in P. leiognathi (Baldwin et al. 1979; O’Kane and Prasher 1992; Meighen and Dunlap 1993; Ast and Dunlap 2004; Dunlap 2009). Although the evolutionary origin of luxA and other bacterial luminescence genes remains obscure (Dunlap and Kita– Tuskamoto 2006), the conserved gene content and gene order of the lux operon in bacteria, luxCDABEG, and the high levels of lux gene and Lux protein amino acid sequence identities among luminous bacteria (e.g., Meighen and Dunlap 1993) leave little doubt of the homology of all presently known bacterial lux operons. Furthermore, the general congruence of phylogenies based on lux genes and other protein coding genes (and the 16S rRNA gene) (Urbanczyk et al. 2008) suggests that the lux operon is ancestral at least to Aliivibrio, Photobacterium, and Vibrio, and possibly to Vibrionaceae. The association of the fatty acid reductase genes, luxCDE, with luxA might have predated the luxA to luxB gene duplication event. Alternatively, the presence of ERIC sequences flanking luxA and luxB in Ph. 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. The origins and evolution of other luminescence genes are not well understood (O’Kane and Prasher 1992). The evolution of bacterial luminescence system also involved recruitment of regulatory and other genes to the lux operon. The lux operons of certain Aliivibrio species contain two regulatory genes, luxR and luxI (> Fig. 13.3), the protein products of which mediate a population density-responsive autoinduction, that is, quorum sensing. Recruitment of regulatory genes to the lux operon during evolution of Aliivibrio presumably enhanced quorum sensing control of luminescence (Dunlap 2009). Furthermore, as mentioned above, luminous Photobacterium strains carry genes involved in the synthesis of riboflavin, the ribEBHA genes, as part of the lux-rib operon (Lee et al. 1994; Ast et al. 2007b). Recruitment of the rib genes to the lux operon likely happened in an ancestor of Photobacterium, since other luminescent bacteria contain the rib genes elsewhere in the genome and not associated with the lux operon (e.g., Callahan and Dunlap 2000). An interesting exception to that general pattern is the presence of ribB initially named (luxH) as the last gene of the lux operon in V. campbellii (previously classified as V. harveyi; Lin et al. 2010). The production of light consumes a substantial amount of energy, through the synthesis of Lux proteins and through their activity (Dunlap and Greenberg 1991). This energetic cost, which may explain the fact that luciferase synthesis is regulated in most luminous bacteria, suggests that activity of the

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Luminous Bacteria

luminescence system plays an important role in the physiology and ecology of luminous bacteria. Most attention to what that role might be has focused on oxygen. One consideration is that, as noted above, the light-emitting reaction might have arisen evolutionarily as a detoxification mechanism, removing oxygen and thereby allowing an organism that is otherwise anaerobic to survive. Related to this possibility is that luciferase, as an oxidase, might function as a secondary respiratory chain that is active when oxygen or iron levels are too low for the cytoplasmic membrane-associated electron transport system to operate. This activity would allow cells expressing luciferase to reoxidize reduced coenzyme even when oxygen levels are low (Hastings and Nealson 1981; Hastings 1983; Nealson and Hastings 1992). Consistent with this view, growth of cytochrome-deficient luminous bacteria is dependent on induction of luciferase, limitation for iron stimulates light production, low oxygen levels promote the luminescence of some luminous bacteria, and luciferase synthesis can be induced under anaerobic conditions (Eberhard et al. 1979; Haygood and Nealson 1985; Makemson 1986; Makemson and Hastings 1982; Nealson and Hastings 1977, 1979). As an alternative to the electron transport system, the activity of luciferase in reoxidizing reduced coenzyme could permit cells of luminous bacteria in low oxygen habitats, such as in animal gut tracts, to continue to transport and metabolize growth substrates, thereby continuing to gain energy through substrate-level phosphorylation. Furthermore, light production presumably facilitates dissemination of luminous bacteria. The feeding of animals on luminous particles (decaying tissues, fecal pellets, and moribund animals infected by luminous bacteria), to which they are attracted, would bring the bacteria into the animal’s nutrient-rich gut tract for additional rounds of reproduction followed by dispersal (Hastings and Nealson 1981; Nealson and Hastings 1992), and recent evidence supports this possibility (Zarubin et al. 2012). Alternatively, the function of the bacterial lux system might be to generate a halotolerant flavodoxin, with light emission an incidental consequence (Kasai 2006). Future studies may test and possibly provide additional support for these and other proposed functions for luminescence, such as a physiological role for luciferase activity in bioluminescent symbioses, but it is not yet clear what factors, physiological or ecological, actually select for the retention and expression of this energetically expensive enzyme system.

Horizontal Acquisition of the Bacterial lux Genes Inheritance of the lux genes has been shown to be primarily vertical. However, some instances of acquisition by horizontal transfer have been identified (Ast et al. 2007; Urbanczyk et al. 2008). In the instances identified, horizontal acquisition of lux genes within Vibrionaceae has been found to be limited to species within the same genera, and no instance of the horizontally transferred genes replacing vertically inherited lux operons has been reported. In contrast to the proposal that horizontal gene transfer drives bacterial speciation (e.g., Gogarten et al. 2002;

Ochman et al. 2000), horizontal acquisition of lux genes apparently has not led to phylogenetic divergence of the recipients (Urbanczyk et al. 2008). The predominant pattern of vertical inheritance of the lux genes, together with the fact that most species of luminous bacteria are members of Vibrionaceae, leads to the hypothesis that these genes arose in an ancestor of Vibrionaceae. The scattered incidence of luminous members in Vibrionaceae, with many nonluminous species and many species with nonluminous strains, indicates that the lux genes have been lost from many descendants of this putative ancestor (Urbanczyk et al. 2008; Dunlap 2009). In Photobacterium, many strains of P. leiognathi carry two intact and apparently functional lux-rib operons in their genomes (Ast et al. 2007b). This situation represents an unusual case of natural merodiploidy in bacteria, the presence of two or more copies of the same gene or genes in the genome of a bacterium, because of the large number of genes involved and because the second operon did not arise by tandem duplication of the first. The two lux-rib operons are distinct in sequence and genomic location. One operon, lux-rib1, is in the ancestral chromosomal location of the lux-rib operon in P. leiognathi and related bacteria. The other, lux-rib2, is located elsewhere in the genome and is present in many but not all strains of P. leiognathi; it is flanked by genes coding for transposases, which suggests it can transfer between strains. Phylogenetic analysis indicates that the lux-rib1 and lux-rib2 operons are more closely related to each other than either is to the lux and rib genes of other bacterial species (Ast et al. 2007b). This finding rules out interspecies horizontal transfer as the origin of the lux-rib2 operon in P. leiognathi; instead, lux-rib2 apparently arose in the distant past within a lineage of P. leiognathi that either has not yet been sampled or has gone extinct. Merodiploidy of the lux-rib operon in P. leiognathi also is the first instance of merodiploid strains of a bacterium having a nonrandom geographic distribution; strains bearing a single lux-rib operon are found over a wide geographic range, whereas lux-rib merodiploid strains have been found only in coastal waters of Honshu, Japan (Ast et al. 2007b; Urbanczyk et al. 2012b). The presence of multiple copies of each of the lux and rib genes might provide opportunities for sequence divergence and selection that could lead to the evolution of new gene functions in one or the other of the duplicate genes. The P. leiognathi lux-rib2 operon has also been found in two strains of P. mandapamensis, which also carry a normal P. mandapamensis lux-rib operon, and in a strain of P. damselae, a species not previously known to be luminous (Urbanczyk et al. 2008). Furthermore, evidence has been obtained indicating horizontal acquisition of the lux genes by a recently recognized species, P. aquimaris (Yoshizawa et al. 2009b; Urbanczyk et al. 2012a). With respect to S. hanedai and S. woodyi, comparison of genes flanking the lux operons suggested that these species had acquired lux genes from a member of Aliivibrio (Kasai et al. 2007), a possibility confirmed through phylogenetic analysis (Urbanczyk et al. 2008). In Photorhabdus species as well, the

Luminous Bacteria

luxCDABE genes may have been acquired by horizontal gene transfer (Forst et al. 1997), possibly from an ancestor of V. harveyi (Meighen 1999). Differences between ecologically distinct strains of Ph. luminescens in the DNA flanking the lux operon (Meighen and Szittner 1992) raise the possibility that lateral transfer of the lux genes to this species occurred more than once (Forst et al. 1997). However, phylogenetic analysis of the Photorhabdus lux genes in the context of Vibrionaceae and Shewanellaceae sequences did not find support either for or against horizontal acquisition of the lux genes by Ph. luminescens (Urbanczyk et al. 2008). It is possible that substantial sequence divergence of the lux genes has occurred since their transfer to Ph. luminescens, thereby making problematic the identification of their source species. Instances of the horizontal acquisition of lux genes have been identified also in Vibrio (Urbanczyk et al. 2008). The only known luminous strain of the human pathogen V. vulnificus (Oliver et al. 1986) apparently acquired its lux genes from V. harveyi, and in V. chagasii, a species not previously known to be luminous (Thompson et al. 2003), two luminous strains were identified and through phylogenetic analysis were shown to have acquired their lux genes apparently from V. harveyi and V. splendidus, respectively (Urbanczyk et al. 2008). A mechanism for these transfers, however, has not been proposed. It should be noted that most species of Vibrionaceae lack the lux genes and therefore are nonluminous. Also, most strains of some luminous species, such as V. cholerae, are nonluminous. The low incidence of luminous species in the family suggests that the lux genes have been lost over evolutionary time from many of the lineages that have given rise to extant species. It also seems likely that nonluminous variants of luminous species can arise frequently through loss of one of more the core genes of the lux operon, luxCDABE (e.g., Wollenberg et al. 2011). The scattered incidence of lux genes in Vibrionaceae presumably relates to different ecologies of the different species. It is not clear, however, how having and expressing lux genes contributes to the lifestyle of most luminous bacteria, because there are no obvious ecological differences between luminous and nonluminous species except in the case of those species that are bioluminescent symbionts of fish and squids.

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sediment, and suspended particulates from a wide variety of locations (Baumann and Baumann 1981; Harvey 1952; Zobell 1946). They also commonly colonize marine animals as saprophytes, commensal enteric symbionts, and parasites (Baumann and Baumann 1981; Harvey 1952; Kozukue 1952; Makemson et al. 1997; Makemson and Hermosa 1999; Meighen and Dunlap 1993; Ruby and Morin 1979; ZoBell 1946). They can also be isolated from inanimate surfaces and macroalgae (Makemson et al. 1992). A few species of luminous bacteria establish bioluminescent symbiosis with marine fish and squids (Dunlap 2009; Dunlap et al. 2007; Hastings and Nealson 1981; Haygood 1993; Ruby 1996; Ruby and Morin 1978; Visick and Ruby 2006; Urbanczyk et al. 2011a, b). In seawater, the incidence of luminous bacteria generally is low (from 0.01–40 cells per ml; Nealson and Hastings 1992), with higher numbers in coastal seawater and lower numbers in open ocean and deeper waters (Ruby and Nealson 1978; Ruby et al. 1980). Possibly reflecting this variation, metagenomic analyses of different marine waters have identified the presence of genes related to luxA (Martı´n Cuadrado et al. 2007) and conversely showed an absence of bacterial lux genes (Nealson and Venter 2007). Therefore, the geographic distribution of luminous bacteria in the plankton varies substantially. In contrast to their generally low incidence in seawater, luminous bacteria can attain very high numbers in saprophytic, commensal, parasitic, and symbiotic associations with animals (up to 1011 cells per ml in symbiotic habitats; Ruby and Nealson 1976; Dunlap 1984; Nealson and Hastings 1992; Visick and Ruby 2006). For example, luminous bacteria can be readily isolated by enrichment from the muscle tissue and skin of marine fish (e.g., Budsberg et al. 2003; Ast and Dunlap 2005) (> Fig. 13.5), and Photobacterium iliopiscarium, a nonluminous species closely

Habitats and Ecology of Luminous Bacteria The luminous Aliivibrio, Photobacterium, Vibrio, and Shewanella species occur in the marine environment, whereas Photorhabdus species are terrestrial. Vibrio cholerae also occurs in brackish environments and freshwater, although strains of this species also commonly occur in coastal seawater (e.g., Kaeding et al. 2007; Urbanczyk et al. 2008).

Marine Luminous bacteria are globally distributed in the marine environment (> Table 13.1) and have been isolated from seawater,

. Fig. 13.5 Saprophytic growth of luminous bacteria. Luminous bacteria have colonized this slice of fish meat, which was photographed in the dark by the light the bacteria produce. Growth of luminous bacteria in and on surfaces of animal tissues is common in nature. This attribute is one means by which luminous bacteria from the environment can be enriched for and isolated

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related to P. phosphoreum and P. kishitanii, has been isolated from the intestines of several species of cold-water fish and from spoiled packaged fish (Ast and Dunlap 2005; Flodgaard et al. 2005; Onarheim et al. 1994; Urakawa et al. 1999). Saprophytic, commensal, parasitic, and symbiotic habitats have the potential 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; Visick and Ruby 2006), which in turn presumably serve as environmental sources of these bacteria for recolonization of animals. As commensal enteric symbionts of fish, luminous bacteria may contribute significantly to the digestion of crustacean prey through the activity of chitinase (Spencer 1961; Baumann and Schubert 1984; Ramesh and Venugopalan 1989). It should be noted that luminous bacteria coexist with and presumably carry out metabolic activities similar to nonluminous bacteria in these different habitats. Luminous bacteria in general, however, show little specificity when forming opportunistic saprophytic and enteric associations with marine animals such as mussels and clams. This lack of specificity can be attributed to the steady influx of bacteria from the water column, which presumably would prevent selection for specialization (Preheim et al. 2011). The exception to this general lack of specificity is bioluminescent symbiosis, in which the luminous bacteria able to colonize this kind of habitat typically are present as single species. In contrast to their associations with marine animals, luminous bacteria apparently do not commonly colonize the surfaces of marine algae. Agar digestion is often observed among nonluminous Vibrio species and other marine bacteria (e.g., Humm 1946), and various attempts, successful (Makemson et al. 1992) and otherwise, have been made to isolate lightemitting bacteria from algal surfaces. To date, however, only one luminous strain, provisionally identified as a member of V. harveyi, that has the ability to digest agar has been isolated from algae (Fukasawa et al. 1987). The uniqueness suggests that the single known isolate of agar-digesting luminous bacteria might have acquired either the genes for agar digestion or the lux genes by horizontal gene transfer. The distributions and numbers of individual species of luminous bacteria tend to correlate with certain environmental factors (Baumann and Baumann 1981). Primary among these factors are temperature and depth (Ruby and Nealson 1978; Yetinson and Shilo 1979; Ruby et al. 1980; Ramaiah and Chandramohan 1987), salinity (Yetinson and Shilo 1979; Feldman and Buck 1984), and nutrient limitation and sensitivity to photooxidation (Shilo and Yetinson 1980; Makemson and Hastings 1982; Haygood and Nealson 1985a). Temperature, along with being an important environmental factor, can influence whether luminous bacteria from environmental samples are detected. For example, Shewanella hanedai, which is psychrotrophic, grows and produces light at low temperature (e.g., 4–15  C) and grows but does not produce light at room temperature (24  C). Therefore, incubation of platings of environmental samples at lower temperatures may reveal the

presence of other luminous species with naturally temperaturesensitive luminescence systems. 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), a species closely related to S. hanedai, grows and produces light at room temperature. Studies of the distribution and density of luminous bacteria in the marine environment traditionally have used visual detection of luminescent colonies arising from seawater spread on nutrient-containing agar plates to identify the presence of these bacteria. However, there are several kinds of luminous bacteria that can be missed with this method. One kind is bacteria that are physiologically cryptic for luminescence, producing visible light in culture only in response to the addition of inducers or other substances to the growth medium (Boettcher and Ruby 1990; Fidopiastis et al. 1999; Nelson et al. 2007) or that require growth at lower than typical room temperatures for light production. Another kind is bacteria with incomplete or defective lux operons (O’Grady and Wimpee 2008). Furthermore, enzyme assay and antibody methods have detected luciferase in several Vibrio spp. that do not produce visible light in culture (Nealson and Walton 1978; Makemson and Hastings 1986; Kou and Makemson 1988). Similarly, luxA-based DNA probes and PCR amplification of lux gene sequences have identified lux genecontaining bacteria from seawater that do not produce light in culture (Potrikus et al. 1984; Palmer and Colwell 1991; Lee and Ruby 1992; Wimpee et al. 1991; Ramaiah et al. 2000; Grim et al. 2008). These studies demonstrate that bacteria carrying the lux genes are more abundant in the marine environment and more phylogenetically diverse than is revealed by analysis of strains isolated on the basis of the production of readily visible levels of light. A counterpoint to this view, however, is the apparently low incidence of lux gene sequences in metagenomic databases (Martı´n Cuadrado et al. 2007; Nealson and Venter 2007), which suggests that luminous Photobacterium, Vibrio, and Aliivibrio, and presumably nonluminous members of these genera as well, represent a very small fraction of the microscopic plankton.

Freshwater Luminous strains of V. cholerae can be isolated from freshwater and brackish estuarine waters (Desmarchelier and Reichelt 1981; West and Lee 1982; West et al. 1983; Palmer and Colwell 1991; Ramaiah et al. 2000; > Table 13.1), as well as from coastal seawater (e.g., Urbanczyk et al. 2008). The first such strain, isolated in 1893 by F. Kutscher from the Elbe River in Germany (Harvey 1952), was named ‘‘Vibrio albensis’’ and later was synonymized with V. cholerae (Reichelt et al. 1976). This species also infects freshwater crustaceans; 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 strongly luminous specimens of the freshwater

Luminous Bacteria

shrimp, Xiphocaridina compressa, in Lake Suwa, Japan. Initially characterized as Microspira phosphoreum, the bacterium was later redescribed as Vibrio yasakii (Majima 1931). A bacterium responsible for this ‘‘light disease of shrimp’’ was isolated more recently from freshwater shrimp in Lake Biwa, Japan, and identified as non-O1 V. cholerae (Shimada et al. 1995). In addition to V. cholerae in freshwater habitats, strains of P. phosphoreum have been isolated from migrating salmon in the Yukon River, Alaska (Budsberg et al. 2003; Ast and Dunlap 2005); presumably, their association with fish slime protected these marine bacteria from osmotic lysis.

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, Ph. luminescens, Ph. temperata, and Ph. asymbiotica (Fischer–Le Saux et al. 1999; > Table 13.1). Two of the three Photorhabdus species occur as the mutualistic symbionts of soil nematodes of the family Heterorhabditidae (> Table 13.1) (Akhurst and Dunphy 1993; Forst and Nealson 1996; Forst et al. 1997; Gerrard et al. 2006; Kuwata et al. 2008; Waterfield et al. 2009). 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 hemocoel, 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 and combat scavenging organisms, such as nematodes and amoeba (Akhurst 1982; Sicard et al. 2007; Waterfield et al. 2009). 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

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the bacteria in their intestines. Cells of Ph. 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 in Ph. luminescens, which is stimulated in laboratory culture by exogenous aldehyde, is not required for successful symbiosis with the nematode; not all strains of Ph. luminescens produce luminescence (Akhurst and Boemare 1986; Forst and Nealson 1996; Schmidt et al. 1989). 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 in this regard because luminescence apparently was a sign that the wound would heal well (Harvey 1957). Indeed, luminous bacteria will grow and produce light on living mammalian tissue (Johnson 1988). Perhaps antibiotic-producing, nonpathogenic Photorhabdus strains 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 Ph. luminescens and P. temperata (Fischer Le Saux et al. 1999; Tailliez et al. 2010) 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 species, and Haneda cultured the bacteria in media containing 0.5 % salt. However, whether these bacteria were terrestrial (i.e., Photorhabdus), from brackish water (i.e., V. cholerae), or marine in origin 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

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Luminous Bacteria

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 1889; Giard and Billet 1889; Inman 1926). Luminous bacteria inhabit the gut tract and colonize external surfaces of marine crustaceans (Inman 1926; Baross et al. 1978; O’Brien and Sizemore 1979; Lavilla–Pitogo et al. 1992); many are chitinolytic (Spencer 1961; Baumann and Schubert 1984). The bacteria enter the hemocoel 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 Aliivibrio, Photobacterium, and Vibrio (Hastings and Nealson 1981; P. Dunlap, unpubl. data). Nonluminous bacteria undoubtedly cause similar infections that go unnoticed due to the lack of light production. As opportunistic pathogens of marine crustaceans, luminous bacteria and their nonluminous relatives have had a profoundly deleterious effect on commercial prawn mariculture (Owens and Busico–Salcedo 2006; Haldar et al. 2011). 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 growout 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 Pen˜a 1998). Primarily responsible are strains of V. harveyi, though other luminous and nonluminous Vibrio species have been identified (Lavilla–Pitogo et al. 1990; Karunasagar et al. 1994; Lavilla– Pitogo and de la Pen˜a 1998; Suwanto et al. 1998; Leano et al. 1998; Austin and Zhang 2006).

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 identified in fish disease, and recently, A. salmonicida (a pathogen of salmonids and cod) has been shown to contain a lux operon (Nelson et al. 2007). Clinical strains of V. vulnificus and V. cholerae typically are nonluminous, but a luminous strain of V. vulnificus has been isolated from a lethal human infection (Oliver et al. 1986; Kaeding et al. 2007), 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 pigeon’s blood (Harvey 1952). 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). However, O1 or O139 serotypes of V. cholerae, which are responsible for life-threatening cases of human diarrheal disease, do not include the light-producing or luxA gene–containing strains (Palmer and Colwell 1991; Ramaiah et al. 2000; Grim et al. 2008).

Bioluminescent Symbiosis A special attribute of a few of the luminous bacteria is the ability to form highly specific, luminescence-based mutualisms, called bioluminescent symbiosis, with certain marine fish and squids (> Table 13.2). Early work is reviewed in detail by Harvey (1952), Buchner (1965), Herring and Morin (1978), and Hastings and Nealson (1981). In these associations, the animal cultures a dense population of luminous bacteria in a tissue complex called a light organ, providing them with nutrients and oxygen for reproduction and light production. The animal in turn uses the bacterial light for luminescence displays associated with sex-specific signaling, predator avoidance, seeing and attracting prey, or schooling. In most of the bacterially bioluminescent fish, the light organs are associated with the gastrointestinal tract; in others, they are subocular (anomalopids), mandibular (monocentrids), or escal (ceratioids). In squids, the bacterial light organs are found as bilobed structures within the mantle cavity, associated with the ink sac. Accessory tissues associated with the light organ, that is, shutter, lens, and reflector, direct and focus the light the bacteria produce. The light organs open to the external environment, either directly or via the intestinal tract or mantle cavity, allowing the excess bacterial cells to be released from the animal’s light organ into the environment as the light-organ population reproduces. In the cases studied, the members of each new host generation of the animal acquire their symbiotic bacteria from the environment. These associations typically are highly specific at the animal family– bacterial species level; members of a family of fish or squid often all harbor the same individual bacterial species as their symbiont (Harvey 1922, 1952; Okada 1926; Harms 1928; Kishitani 1930; Yasaki 1928; Haneda 1938, 1950; Ahrens 1965; Buchner 1965; Hastings 1971; Morin et al. 1975; Herring 1977; Herring and Morin 1978; Nealson 1979; McFall–Ngai 1983; McFall–Ngai and Dunlap 1983; Haygood et al. 1984; Nealson et al. 1984; Dunlap and McFall–Ngai 1987; Wei and Young 1989; McFall Ngai and Morin 1991; McFall Ngai and Ruby 1991; Ruby and Asato 1993; Graf and Ruby 1998; Wada et al. 1999; Woodland et al. 2002; Sasaki et al. 2003; Jones and Nishiguchi 2004; Sparks et al. 2005; Dunlap et al. 2009; Charkrabarty et al. 2011; Dunlap and Nakamura 2011). The bacteria are housed extracellularly, and in most cases they are known to not be obligately dependent on the host for their reproduction, as they colonize a variety of other habitats (Baumann and Baumann 1981; Hastings and Nealson 1981; Visick and Ruby 2006). Bioluminescent symbiosis

Sepiolidae

+ + + + +

Himantolophidae

Linophrynidae

Melanocetidae

Oneirodidae

Thaumatichtyidae

+

Not identified

+

Candidatus Photodesmus katoptron

+

+

+

Photobacterium mandapamensis

Gigantactinidae

+

+

+

+

Photobacterium kishitanii

+

+

Photobacterium leiognathi

Diceratiidae

+

Aliivibrio wodanis

Ceratiidae

+

Aliivibro ‘‘thorii’’

+

+

+

+

Aliivibrio fischeri

Centrophrynidae

Lophiiformes

Moridae

Merluciidae

Macrouridae

Gadiformes

Chlorophthalmidae

Aulopiformes

Opisthoproctidae

Argentiniformes

Congridae

Anguilliformes

Fish

Sepiolida

Loliginidae

Teuthida

Squids

Host animala

. Table 13.2 Bacterial species affiliations in bioluminescent symbiosis

Luminous Bacteria

13 509

+

+

Photobacterium kishitanii

+

Photobacterium mandapamensis +

Candidatus Photodesmus katoptron

Not identified

Data are from (Ast and Dunlap 2005; Ast et al. 2009; Castle and Paxton 1984; Dunlap et al. 2004; Dunlap et al. 2007; Fidopiastis et al.1998; Fukasawa and Dunlap 1986; Haygood and Distel 1993; Haygood et al. 1992; Hendry and Dunlap 2011; Kaeding et al. 2007; Nishiguchi 2000; Wada et al. 2006; Wolfe and Haygood 1991; and Dunlap unpubl. data.

a

+

+

Photobacterium leiognathi

+

Aliivibrio wodanis

+

Aliivibro ‘‘thorii’’

Leiognathidae

+

Aliivibrio fischeri

Apogonidae

Acropomatidae

Perciformes

Trachichthyidae

Monocentridae

Anomalopidae

Beryciformes

Host animala

13

. Table 13.2 (continued)

510 Luminous Bacteria

Luminous Bacteria

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 bacterially produced nutrient or enzymatic activity needed for host nutrition (Claes and Dunlap 2000). Luminous bacteria might also form symbioses with pyrosomes and salps; little is known, however, and the topic remains controversial (Harvey 1952; Buchner 1965). Pyrosome zooids bear a pair of simple photophores that contain intracellular bacteroids, but the involvement of the bacteroids in pyrosome luminescence has been both discounted and supported (Galt 1978; Herring 1978; Mackie and Bone 1978; Haygood 1993). 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). A similar proposal for luminous myctophid and stomiiform fish, that the luminescence of the fish’s photophores is produced by symbiotic luminous bacteria (Foran 1991), however, has been conclusively refuted (Haygood et al. 1994). The following information focuses primarily, although not exclusively, on bioluminescent symbiosis in fish. Detailed information on the bioluminescent mutualism of A. fischeri with the sepiolid squid Euprymna scolopes can be found in the chapter by K. Visick (Chap. 20, ‘‘Vibrio fisheri: Squid Symbiosis,’’ Vol. 1).

Patterns of Host Affiliation Six species of luminous bacteria form bioluminescent symbioses with fish and squids, A. fischeri, A. ‘‘thorii,’’ A. wodanis, P. kishitanii, P. leiognathi, and P. mandapamensis. Their currently known host affiliations are listed in > Table 13.2. There are over 460 species of bacterially luminous marine fish, in 21 families of seven teleost orders, and several species of squid in two families of two cephalopod orders (Dunlap et al. 2007; Herring and Morin 1978; Nelson 2006) (> Fig. 13.6). The most numerous of these symbiotic bacteria, due to the exceptional abundance of their host animals in the marine environment, are likely to be P. kishitanii and P. leiognathi. The hosts of P. kishitanii are fish of diverse families in deep-sea habitats worldwide, many of which are abundant, and the hosts of P. leiognathi are primarily fish of the family Leiognathidae, which are abundant in shallow coastal waters of Southeast Asia and South Asia (Tiews and Caces Borja 1965; Ku¨hlmorgen–Hille 1974; Herring and Morin 1978; Cohen et al. 1990; Orlov and Iwamoto 2006; Dunlap et al. 2007; Dunlap et al. 2009). The bioluminescent symbionts of deep-sea fish previously were thought to be P. phosphoreum, but detailed phylogenetic analyses of the phosphoreum species group identified P. kishitanii as the species occurring in light organs of deepsea fish. Despite extensive testing, no bonafide member of P. phosphoreum has been found in light-organ symbiosis (Ast and Dunlap 2005; Ast et al. 2009; Dunlap and Ast 2005; Dunlap et al. 2007; Kaeding et al. 2007; Urbanczyk et al. 2007). Similarly, strains of bacteria from light organs of the sepiolid squids Sepiola affinis and Sepiola robusta, previously identified as

13

A. logei (V. logei) (Fidopiastis et al. 1998), were recently identified based on detailed phylogenetic criteria as three entities, A. fischeri; A. ‘‘thorii’’, a newly recognized bacterial clade; and A. wodanis, a previously described species newly recognized as a bioluminescent symbiont; apparently no bonafide member of A. logei has been found in light-organ symbiosis (Ast et al. 2009). In addition to these bacteria, strains identified as V. harveyi have been found in the light organ of larval leiognathid fish (Dunlap et al. 2008); it is not yet known if V. harveyi is present as an incidental, transient colonizer of the nascent light organ of this fish, as a pathogen, or, possibly, as an actual symbiont (Dunlap et al. 2008).

Species Specificity, Cosymbiosis, and Symbiont: Host Codivergence Previously, bioluminescent symbioses were characterized as species specific, with the light organ of each animal thought to harbor a single, pure culture of bacteria and with the members of each family of fish or squids thought to all harbor the same bacterial species as their symbiont (Hastings and Nealson 1981; Nealson and Hastings 1992; Dunlap and Kita–Tsukamoto 2006). This pattern of specificity still generally holds, but several deviations from a strict host family–bacterial species specificity have been identified. On the one hand, individual light organs of certain squid and fish have been found to harbor two bacterial species, a situation termed cosymbiosis. In contradicting a strict one-to-one relationship, cosymbiosis requires the mechanism by which the host might select its symbiotic bacteria, such as surface recognition, to respond to features common to both bacterial species or to distinct features of each (Ast et al. 2009; Fidopiastis et al. 1998; Dunlap et al. 2007; Dunlap et al. 2008; Kaeding et al. 2007; Dunlap et al., unpubl. data). On the other hand, different host species and genera within a family have been found to harbor different species of bacteria (> Table 13.2). One example of this breakdown of host family level bacterial specificity is the presence in light organs of Acropoma hanedai of P. kishitanii, whereas P. mandapamensis is present as the primary symbiont in light organs of Acropoma japonicum. Another is the presence in different species of Coelorinchus of P. kishitanii or A. fischeri (Dunlap et al. 2007; Kaeding et al. 2007; Wada et al. 2006). These discrepancies suggest that a strict genetically based host selection of a specific symbiotic bacterium (McFall Ngai and Morin 1991) may not be operative in bioluminescent symbiosis or may not be operative for all bacterially luminous animals. Consistent with this possibility is the lack of codivergence, that is, cospeciation, between host and symbiont lineages (> Fig. 13.7). Genetic selection might reasonably lead to a codivergence, as reported for squid-symbiotic bacteria (Nishiguchi et al. 1998), but instead of congruent host and symbiont phylogenies, detailed phylogenetic analysis based on homologous genes, suitable numbers of strains, and a diversity of hosts reveals instead that the patterns of symbiont affiliations for fish and squids are strikingly noncongruent

511

512

13

Luminous Bacteria

. Fig. 13.6 Bacterially luminous fish. Shown are a few of the more than 460 species of fish that host luminous bacteria as bioluminescent symbionts. Counterclockwise from the top, the fish are Physiculus japonicus (Gadiformes: Moridae) (photo by A. Fukui), host of P. kishitanii and, less commonly, A. fischeri; Eubleekeria jonesi (Perciformes: Leiognathidae) (photo by P.V. Dunlap), host of P. leiognathi; Acropoma japonicus (Perciformes: Acropomatidae) (photo by A. Fukui), host of P. mandapamensis and, less commonly, P. leiognathi; Chlorophthalmus nigromarginatus (Aulopiformes: Chlorophthalmidae), host of P. kishitanii; Monocentris japonicus (Beryciformes: Monocentridae) (photo by P.V. Dunlap), host of A. fischeri; and Aulotrachichthys prosthemius (Beryciformes: Trachichthyidae) (photo by A. Fukui), host of P. kishitanii

(Dunlap et al. 2007). Furthermore, phylogenetically distantly related hosts, for example, bacterially luminous aulopiforms, most gadiforms, and certain beryciforms, all harbor the same bacterial species, P. kishitanii, whereas some closely related hosts, such as the acropomatid fish A. hanedai and A. japonicum, as noted above, harbor distinct species, P. kishitanii and P. mandapamensis, respectively (Dunlap et al. 2007) (> Fig. 13.7).

An alternative hypothesis to account for the observed patterns of symbiont–host affiliation in bioluminescent symbiosis is environmental congruence. This hypothesis, first outlined by Hastings and Nealson (1981), links the differing environmental distributions of different species of luminous bacteria, that is, where each species is most abundant, with the environmental distribution of its host animal (Dunlap et al. 2007; Hastings and

Luminous Bacteria Loliginidae

Aliivibrio fischeri

Squid Sepiolidae

Aliivibrio “thorii ” Aliivibrio wodanis

Opisthoproctidae C hlorophthalmidae

Photobacterium leiognathi

Macrouridae Merluciidae

Photobacterium kishitanii

Moridae

Fish

Anomalopidae

Photobacterium mandapamensis

Monocentridae Candidatus Photodesmus katoptron

Trachichthyidae Acropomatidae Apogonidae

?

Vibrio harveyi

Leiognathidae

. Fig. 13.7 Host affiliations of symbiotic luminous bacteria. Families of bacterially luminous squids and fish are listed on the left, with lines to the corresponding bacterial species on the right that have been isolated from light organs of these animals. Different members of individual families of fish and some squids often harbor different species of bacteria, in some cases within the light organ of the same host specimen. Some of the bacteria, for example, A. fischeri, P. kishitanii, P. mandapamensis, are found in light organs of a diversity of fish and squids. These attributes highlight the lack of strict family level bacterial species specificity and the lack of phylogenetic congruence between host and symbiont in bioluminescent symbiosis (Dunlap et al. 2007; Kaeding et al. 2007). The question mark for the link from Leiognathidae to V. harveyi reflects the single instance that this bacterial species has been isolated from light organ symbiosis (Dunlap et al. 2008)

Nealson 1981; Kaeding et al. 2007; Dunlap et al. 2008). Temperature, which influences the presence and relative numbers of the different species of luminous bacteria in the marine environment, may be the key environmental factor; deeper, colder dwelling hosts harbor the more psychrotrophic luminous species found in those habitats, P. kishitanii, for example, as their bioluminescent symbiont, whereas shallower and warmer dwelling hosts harbor the more mesophilic luminous species found in those habitats, A. fischeri and P. leiognathi, for example. An important further consideration is the ontogenetic ecology of the host. Early life history stages of these animals, for example, eggs, larvae, and juveniles, often are distributed in the environment differently from adults. The key factor therefore may be where in the environment the animal is when it is developmentally ready to initiate symbiosis. The luminous bacterial species most abundant in and adapted to the conditions of those habitats presumably would the ones most likely to initiate symbiosis (Dunlap et al. 2007; Kaeding et al. 2007). Information about

13

early life history stages of bacterially luminous animals, especially fish, is very limited, but evidence is beginning to accumulate that supports the environmental congruence hypothesis (Dunlap et al. 2008). Nonetheless, some form of host selection must be occurring because to date only luminous bacteria, and only certain species of luminous bacteria, have been found in light organs of fish and squid. Most likely, a combination of environmental congruence and some level of selection are operative. The luminous bacteria symbiotic with two other groups of fish, the flashlight fish, family Anomalopidae, and bacterially luminous deep-sea anglerfish, in order Lophiiformes, present a possible contrast to the apparent lack of strict species specificity in bioluminescent symbiosis. Microscopic analysis showing the presence of masses of bacterial cells within the light organs, assays specific for bacterial luciferase, and other studies convincingly demonstrate the bacterial nature of light emission in these fish (Bassot 1966; Harvey 1922; Haygood et al. 1984; Kessel 1977; Leisman et al. 1980; Munk et al. 1998), but the bacteria from light organs of these fish have not been grown in laboratory culture despite numerous attempts (Hastings and Nealson 1981; Haygood 1993; Hendry and Dunlap 2011; Herring and Morin 1978). The inability to grow these bacteria in the laboratory suggests that they have lost the ability to reproduce outside the host light organ and are therefore might be obligately dependent on their hosts (Haygood 1993; Haygood and Distel 1993). An obligate relationship presumably would lead to a very high degree of specificity and possibly also to codivergence between the host fish and its bacteria. 16S rRNA gene sequence analysis of the symbiotic bacteria of two ceratioids, representing different families of lophiiformes, places these bacteria as members of Vibrionaceae and possibly as a new bacterial species in each fish (Haygood and Distel 1993; Hendry and Dunlap 2011). Analysis of the luxA and 16S rRNA genes of anomalopid symbionts suggested these bacteria are members of Vibrio and that different genera of the fish harbor bacteria that differ at greater than the strain level (Haygood 1990; Wolfe and Haygood 1991). Consistent with and extending these findings, a recent detailed multilocus analysis classified the bacteria symbiotic with the anomalopid fish Anomalops katoptron as a new Vibrionaceae genus and species, Candidatus Photodesmus katoptron, which is closely related to Vibrio (Hendry and Dunlap 2011).

Symbiont Acquisition In the few cases studied, bacterially luminous squid and fish have been found to acquire their symbiotic luminous bacteria from the environment with each new host generation. The sepiolid squid E. scolopes acquires cells of A. fischeri from seawater shortly after hatching from the egg as aposymbiotic juveniles; bacteria other than the native symbiont fail to colonize the light organ or do so less effectively (Wei and Young 1989; McFall Ngai and Ruby 1991). In fish, the symbiotic bacteria are acquired later, following development of larvae and inception of light organ formation (Wada et al. 1999; Dunlap et al. 2008;

513

514

13

Luminous Bacteria

. Fig. 13.8 Developing light organ. This electron micrograph of a section of the light organ of the fish Nuchequula nuchalis (Leiognathidae) (micrograph prepared by Sasha Meschinchi, Microscopy and Imaging Laboratory, University of Michigan) shows tubules of the nascent light organ. Some tubules are empty, whereas some are filled or becoming filled with bacteria

Dunlap et al. 2009; Dunlap et al. 2012) (> Fig. 13.8), which is consistent with acquisition of the bacteria from the environment. Also consistent with environmental acquisition, symbiotic bacteria apparently are not present on eggs within the ovary of anomalopid fish (Haygood 1993).

Quorum Sensing Control (Autoinduction) of Bacterial Luminescence In many luminous bacteria, luciferase synthesis and luminescence are regulated in a population density-responsive manner. At low population density, very little luciferase is synthesized, and consequently, little light is produced, whereas at high population density, luciferase levels are induced 100–1,000-fold and light levels increase by 103–106-fold. Population density–responsive induction of luciferase synthesis and luminescence is controlled in part by the production and accumulation in the cell’s local environment of small signal molecules, called autoinducers (acyl-homoserine lactones, AHLs, and other low molecular weight compounds), which function via regulatory proteins to activate transcription of the lux operon. Originally called autoinduction and discovered through study of the pattern of luminescence and luciferase production of V. harveyi in batch culture (Nealson et al. 1970), this gene regulatory mechanism is now referred to as quorum sensing to reflect its relationship with

population density (Fuqua et al. 1994; Greenberg 1997; Hastings and Greenberg 1999; Miller and Bassler 2001). Over the past 30 years, there has been a very substantial accumulation of information on how two luminous bacteria, V. harveyi (a key strain recently was reclassified as V. campbellii; Lin et al. 2010) and A. fischeri, regulate luminescence by quorum sensing. Studies in these two bacteria established a base of knowledge that led to the discovery of biochemically and genetically homologous quorum sensing systems in a wide variety of nonluminous bacteria; quorum sensing controls many cellular activities other than light production, particularly the production of extracellular enzymes and other extracellular factors thought to be adaptive for bacteria at high population density and in association with animal and plant hosts (Fuqua et al. 1996; Dunlap 1997; Bassler, et al. 1997; Swift et al. 1999; Callahan and Dunlap 2000; Waters and Bassler 2005; Dunlap and Kita– Tuskamoto 2006; Higgins et al. 2007). Quorum sensing therefore is not only not unique or even special to luminous bacteria, it is also widespread and evolutionarily conserved across a diversity of bacteria. As a signal–response mechanism by which bacteria can assess their local population density, quorum sensing might have arisen evolutionarily as a diffusion sensor or efficiency sensor (Redfield 2002; Hense et al. 2007), mediating whether or not cells produce extracellular enzymes and other factors for obtaining nutrients. Substantial attention has been placed recently on the definition and correct usage of terms for quorum sensing and other chemically mediated bacterial interactions (Platt and Fuqua 2010; Stacy et al. 2012). This form of genetic regulation has been studied in detail in A. fischeri and V. harveyi but remains poorly understood in other luminous bacteria (Meighen 1999). It should be noted also that some luminous bacteria express luminescence constitutively during growth in batch culture (Katznelson and Ulitzur 1977; P. Dunlap, unpubl. data) and therefore apparently do not use a quorum sensing system to control luminescence. An overview of quorum sensing in V. harveyi and A. fischeri is provided here, and more information on this topic is provided in the chapters by B. Bassler (Chap. 22, ‘‘Quorum Sensing,’’ Vol. 2) and K. Visick (Chap. 20, ‘‘Vibrio fisheri: Squid Symbiosis,’’ Vol. 1). Early Studies of Quorum Sensing Control of Luminescence. In A. fischeri and V. harveyi, expression of the lux operon, which initially is 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). Early analyses of the ‘‘phases of luminescence’’ in culture (e.g., 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 A. fischeri and V. harveyi were found also to release into the medium species-specific positively acting secondary metabolites, called autoinducers. These compounds accumulate in the growth medium in a population densitydependent manner, and once they attain threshold concentrations, they induce luciferase synthesis (Nealson et al. 1970;

Luminous Bacteria

Eberhard 1972; Nealson 1977; Nealson and Hastings 1979; Ulitzur and Hastings 1979; Rosson and Nealson 1981). Analysis of quorum sensing attained benchmarks of progress in the 1980s with the identification of autoinducer signal molecules and lux regulatory genes. The first autoinducer, 3-oxohexanoyl-HSL (3-oxo-C6-HSL), and the first lux regulatory genes, luxI (encoding 3-oxo-C6-HSL synthase; Schaefer et al. 1996) and luxRAf (encoding acyl-HSL receptor/transcriptional activator), were identified in A. fischeri (Eberhard et al. 1981; Engebrecht et al. 1983; Engebrecht and Silverman 1984), followed by identification of 3-hydroxybutanoyl-HSL and a nonhomologous regulatory gene, luxRVh, in V. harveyi (Cao and Meighen 1989; Martin et al. 1989; Showalter et al. 1990). Quorum Sensing Regulatory Circuitry. Ongoing progress since the 1980s has substantially deepened understanding of the quorum sensing genetic regulatory circuitry controlling luminescence in V. harveyi and A. fischeri. The two regulatory

C AI-1

13

systems are strikingly different. In V. campbellii (previously classified as V. harveyi; Lin et al. 2010), three chemically distinct autoinducers are produced, 3-hydroxybutanoyl-HSL (harveyi autoinducer-1, HAI-1), (2 S,4 S)-2-methyl-2,3,3,4tetrahydroxytetrahydrofuran borate (V. harveyi autoinducer-2, and (S)-3-hydroxytridecan-4-one (cholerae AI-2Vh), autoinducer, CAI-1) (Cao and Meighen 1989; Cao and Meighen 1993; Chen et al. 2002; Higgins et al. 2007) (> Fig. 13.9). Synthesis of HAI-1 is dependent on LuxM (Bassler et al. 1993), synthesis of AI-2Vh is catalyzed by LuxS (Schauder et al. 2001), and synthesis of CAI-1 is catalyzed by CqsA (Kelly et al. 2009; Wei et al. 2011). Each of these molecules is specifically recognized by a different cytoplasmic membrane-associated twocomponent histidine kinase receptor, LuxN (Bassler et al. 1993; Freeman et al. 2000), LuxPQ (Bassler et al. 1994b), and CqsS (Henke and Bassler 2004), respectively (> Fig. 13.9). When concentrations of the autoinducers are low, such as at low

HAI-1

AI-2Vh

OM LuxP C M C qsS LuxM

LuxN

LuxQ

LuxS

C qsA

LuxU

LuxO~P σ 54 Qrr1,5 Qrr2,3,4

LuxRVh

other QS-regulated genes

luxCDABEGH

. Fig.13.9 Regulatory circuitry controlling luminescence in V. campbellii. The expression of lux operon, and of other quorum sensing-regulated genes, in V. campbellii (previously classified as V. harveyi), is coordinated by three chemically distinct autoinducers, HAI-1, AI-2Vh, and CAI-1, that modulate the phosphorylation state of luxU. The synthesis of each autoinducer is catalyzed by a different protein, LuxM, LuxS, and CqsA, and each is recognized by a different cytoplasmic membrane-associated two-component histidine kinase receptor, LuxN, LuxPQ, and CqsS, respectively. Low concentrations of the autoinducers lead to phosphorylation of LuxO and via quorum regulatory RNAs to the destabilization the luxRVh transcript, thereby blocking lux operon transcriptional activation by LuxRVh. High concentrations of the autoinducers reverse the phosphorylation cascade, allowing formation of LuxRVh and activation of lux operon transcription. Arrows indicate positive interactions or transcriptional activation, whereas bars indicate blocking of transcription. See the text for details and references (Redrawn from Tu et al. (2010))

515

516

13

Luminous Bacteria

population density or in habitats in which the autoinducers diffuse away rapidly from cells, that is, seawater, the receptor proteins function as kinases, transferring phosphate to LuxU, a histidine-phosphotransfer protein. LuxU then transfers the phosphate to LuxO, a DNA binding response regulator, the expression of which is subject to repression by LuxT (Bassler 1999; Bassler et al. 1994a; Cao et al. 1995; Freeman and Bassler 1999a, b; Surete et al. 1999; Lilley and Bassler 2000; Lin et al. 2000; Miyamoto et al. 2003; Waters and Bassler 2006). LuxO  P, together with sigma factor s54, then activates expression of genes coding for five small quorum regulatory RNAs (Qrr), Qrr1 through Qrr5 (Lenz et al. 2004; Tu and Bassler 2007). The Qrr RNAs bind and destabilize the luxRVh transcript, blocking production of LuxRVh protein, the transcriptional activator of the lux operon (Showalter et al. 1990; Swartzman et al. 1992), and thereby preventing activation of lux operon transcription. Conversely, once autoinducer concentrations have attained high levels in the cell’s local environment, they bind to their receptors, causing the receptors to switch from kinases to phosphatases, leading to the dephosphorylation of LuxO. With the resulting cessation of qrr gene transcription, luxRVh message is produced and translated, and LuxRVh activates lux operon transcription. Negative autoregulation of LuxRVh adds additional complexity to this system (Chatterjee et al. 1996;

Miyamoto et al. 1996), as does the negative autoregulation of LuxO and posttranscriptional control of LuxO by Qrr sRNAs (Tu et al. 2010; > Fig. 13.9). The complexity of this regulatory system apparently benefits V. campbellii by allowing a fine tuning of its quorum sensing response to differences in the various habitats the bacterium colonizes (Waters and Bassler 2005; Ng and Bassler 2009; Tu et al. 2010). Quorum sensing control of luminescence in A. fischeri involves a very different regulatory system. A population density-dependent accumulation of the autoinducer 3-oxohexanoyl-homoserine lactone (3-oxo-C6-HSL), a membranepermeant molecule, triggers induction when it reaches a critical concentration (> Fig. 13.10). Synthesis of 3-oxo-C6HSL is catalyzed by LuxI, an acyl-homoserine lactone synthase. The regulatory genes, luxRAf and luxI, are directly linked to the lux operon (> Figs. 13.2, > 13.10). The luxRAf gene, which is upstream of the lux operon and divergently transcribed from it, encodes a transcriptional activator protein, LuxRAf, which associates with 3-oxo-C6-HSL, forming a complex that binds at a site in the lux operon promoter and that facilitates the binding of RNA polymerase, thereby activating transcription of the genes for light production, luxICDABEG. Because luxI is a gene of the lux operon, increased transcription leads to increased synthesis of 3-oxo-C6-HSL in an autocatalytic, positive feedback manner.

. Fig. 13.10 Regulatory circuitry controlling luminescence in A. fischeri. The expression of the lux operon, and of other quorum sensing-regulated genes, in A. fischeri is mediated primarily by the concentration of AI-1, which forms a complex with LuxRAf. Synthesis of AI-1 is dependent on LuxI, and the AI-1/LuxRAf complex activates luxICDABEG transcription. Together with cAMP, the CRP protein activates expression from the luxRAf promoter, increasing synthesis of LuxRAf and potentiating the system to induce strongly once sufficient AI-1 has accumulated. Increased expression from the lux operon promoter leads to a stimulation of AI-1 synthesis in an autocatalytic, positive feedback manner, resulting in a rapid and strong induction of luciferase synthesis. A second autoinducer, AI-2Af, interacts with LuxRAf, interfering with the interaction between AI-1 and LuxRAf. The hypothesized AI-2Af/LuxRAf complex is thought to be transcriptionally less effective and functions to delay the onset of AI1/LuxRAf activation of luxICDABEG transcription. See the text for details and references. Figure provided by K. Dougan, University of Michigan

Luminous Bacteria

The result is a rapid and strong induction of luciferase synthesis and luminescence (Engebrecht et al. 1983; Engebrecht and Silverman 1984; Kaplan and Greenberg 1985; Eberhard et al. 1991; Schaefer et al. 1996; Stevens and Greenberg 1997). Other regulatory factors modulate quorum sensing in A. fischeri. GroEL is necessary for production of active LuxRAf (Adar et al. 1992; Adar and Ulitzur 1993; Dolan and Greenberg 1992), and 3’ 5’-cyclic AMP (cAMP) and cAMP receptor protein (CRP) activate transcription of luxR and thereby potentiate the cell’s response to 3-oxo-C6-HSL (Dunlap and Greenberg 1985; 1988; Dunlap 1989; Dunlap and Kuo 1992). LuxRAf/3-oxo-C6HSL also negatively autoregulates luxRAf expression (Dunlap and Greenberg 1988; Dunlap and Ray 1989), and a second autoinducer, octanoyl-HSL, synthesis of which is catalyzed by AinS, interacts with LuxRAf apparently to delay lux operon induction (Eberhard et al. 1986; Kuo et al. 1996; Hanzelka et al. 1999) (> Fig. 13.10). Under anaerobic conditions, Fnr contributes to lux operon expression (Mu¨ller Breitkreutz and Winkler 1993). A homolog of the V. harveyi luxO gene is carried by A. fischeri, and as with V. harveyi, LuxO in A. fischeri functions as a repressor of luminescence (Miyamoto et al. 2000, 2003), apparently in a qrr-dependent manner (Miyashiro et al. 2010). In addition, LitR, which has substantial sequence similarity to LuxRAf, positively regulates lux operon expression (Fidopiastis et al. 2002), and LexA is thought to contribute to control of luminescence (Shadel et al. 1990; Ulitzur and Dunlap 1995). Despite the many differences in the quorum sensing regulatory systems of V. harveyi and A. fischeri, there are some commonalities. The C-terminal half of the A. fischeri AinS protein is 34 % identical to the V. harveyi LuxM protein, and the Nterminal portion of A. fischeri AinR, encoded by ainR, a gene downstream of ainS, is 38 % identical to the amino terminus of V. harveyi LuxN (Gilson et al. 1995). Whether AinR itself, possibly through interaction with C8-HSL, plays a role in lux regulation in A. fischeri (Gilson et al. 1995; Kuo et al. 1994) has not been established. The deduced amino acid residue sequence of A. fischeri LuxO is approximately 70 % identical to that of V. harveyi (Miyamoto et al. 2000), and a gene immediately downstream of luxO in A. fischeri is likely to be a homolog of V. harveyi luxU. Whether these homologies indicate overlaps in quorum sensing control, however, remains to be determined. Another commonality between the two systems is expression of the lux operons of both species is dependent on cyclic AMP (cAMP) and cAMP receptor protein (CRP) (Ulitzur and Yashphe 1975; Chen et al. 1985; Dunlap and Greenberg 1985; 1988; Dunlap 1989; Dunlap and Kuo 1992). Consistent with this dependence, the regulatory regions upstream of the lux operons of both species contain a CRP binding site (Engebrecht and Silverman 1987; Devine et al. 1988; Miyamoto et al. 1988). Mutants of V. harveyi and A. 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 1989). Furthermore, CRP from V. harveyi has been purified and shown to be immunologically and functionally homologous to CRP of Escherichia coli

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(Chen et al. 1985), and the cya and crp genes of A. 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., unpubl. data). In V. harveyi, cAMP-dependent binding of CRP activates lux operon expression as well as expression of luxRVh (Chatterjee et al. 2002). Studies with A. fischeri and with E. coli carrying the A. fischeri luxR–luxICDABEG genes indicate that a major effect of cAMP–CRP is to activate expression of LuxRAf while repressing transcription from the lux operon promoter (Dunlap and Greenberg 1985; 1988; Dunlap and Kuo 1992; Shadel et al. 1990), although other important lux regulatory effects have also been described (Shadel and Baldwin 1991; 1992a, b). Control by cAMP–CRP suggests that the luminescence systems of these bacteria might be part of the cellular response to stresses associated with nutrient limitation and decreasing growth rate. Physiological Control of Luminescence. The presence of glucose can suppress bacterial luminescence; this catabolite repression effect presumably operates by modulating the levels of cAMP and CRP in the cell (Nealson et al. 1972; Meighen and Dunlap 1993; Dunlap 1997). 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 A. fischeri is temporary, not reversed 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 A. fischeri grown in phosphate-limited chemostat culture; glucose repression of luminescence under these conditions is permanent, and it is reversible by cAMP (Friedrich and Greenberg 1983). A further complication for studies of cAMP control of luminescence in A. 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. 1992; Dunlap and Callahan 1993; Callahan et al. 1995). In addition to glucose, other physiological factors can strongly influence the amount of light produced by luminous bacteria grown in laboratory culture. Oxygen, amino acids, iron, and osmolarity have distinct effects, depending on the species studied (Harvey 1952; Coffey 1967; Nealson et al. 1970; Hastings and Nealson 1977; Makemson and Hastings 1982; Dunlap 1985; Haygood and Nealson 1985a; Hastings et al. 1987; Guerrero and Makemson 1989; Dunlap 1991). Those factors that stimulate growth rate, such as readily metabolized carbohydrates, tend to decrease light production and luciferase synthesis. They do so presumably by directing the consumption of oxygen and reducing power (FMNH2) away from luciferase (e.g., McElroy and Seliger 1962; Coffey 1967) and by indirectly or directly influencing lux gene expression (Dunlap and Greenberg 1985; Dunlap 2000). Conversely, factors that restrict growth rate, such as limitation for iron and low or high osmolarity of the growth medium, depending on the species, tend to stimulate the synthesis and activity of luciferase (Hastings and Nealson 1977;

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Makemson and Hastings 1982; Haygood and Nealson 1985; Hastings et al. 1987; Dunlap 1991). The mechanisms by which these factors operate, however, are not well understood, although they presumably interface with the quorum sensing control circuitry in some way. Other Genes Subject to Quorum Sensing Regulation. Studies of V. harveyi led to the first demonstration of non-lux quorumsensing-regulated genes. In V. harveyi, the production of the fatty acid storage product poly-b-hydroxybutyrate is controlled in a cell density-dependent manner by 3-OH-C4-HSL (Sun et al. 1994; Miyamoto et al. 1998). Along with that finding, LuxRVh has been shown to control various non-lux genes and to act as either a transcriptional activator or repressor (Chatterjee et al. 1996; Miyamoto et al. 1996; Miyamoto and Meighen 2006; Waters and Bassler 2006; Pompeani et al. 2008). In A. fischeri, proteomic analysis of mutants defective in luxR, luxI, and ainS revealed the presence of several quorum-sensing-controlled non-lux genes, components of a LuxR-dependent quorumsensing regulon; these genes code for a variety of different of proteins, some of which apparently contribute to the ability of A. fischeri to colonize its squid host, E. scolopes (Callahan and Dunlap 2000; Qin et al. 2007). Transcript analysis confirmed and extended these results to several additional genes (Antunes et al. 2007).

Isolation, Cultivation, Storage, and Identification of Luminous Bacteria When working with luminous bacteria, and particularly when isolating new strains from nature, the possibility that these bacteria could be pathogenic (e.g., Kaeding et al. 2007) should always be kept in mind and appropriate care to avoid infection should always be used. Additional and detailed information on the isolation, cultivation, and phenotypic characterization of luminous bacteria can be found in Nealson (1978), Baumann et al. (1984), Baumann and Baumann (1981), Farmer and Hickman–Brenner (1992), and Baumann and Schubert (1984).

Isolation Luminous bacteria can be isolated from most marine environments, and two methods, direct plating of seawater and enrichment from marine fish skin, are effective and simple for this purpose. An easily prepared complete medium that is suitable for growing all known luminous bacteria, LSW-70, contains natural or artificial seawater, diluted to 70 % of full strength, 10 g l–1 tryptone or peptone, and 5 g l–1 yeast extract, with 15 g l–1 agar for solid medium (Dunlap et al. 2004). Sugars and sugar alcohols (i.e., glucose, glycerol) are unnecessary for good growth and luminescence and can lead to acid production and death of cultures (Hill 1928; Johnson and Shunk 1936; Dunlap et al. 1995); their use in isolation media should be avoided. For isolations from environments where high numbers of bacteria that form spreading colonies may be present, such as coastal

seawater, sediment, and intestinal tracts of marine animals, the use of agar at 4 % (40 g l–1) (Baumann et al. 1984) is recommended. This harder, less moist agar limits the ability of bacteria motile on solid surfaces, for example, certain peritrichously flagellated bacteria and bacteria that move by gliding motility, to spread over the plate and cause cross contamination of colonies. Direct plating of seawater involves simply spreading an appropriate volume, typically 10–100 ml for coastal seawater, of the sample on one or more plates and incubating at room temperature or, preferably, cooler temperatures, such as 15–20  C. For open ocean seawater and other samples with a lower number of bacteria, larger volumes, for example, 100 ml to 1 l, can be filtered through membrane filters with a pore size of 0.2 mm or 0.45 mm to collect the bacteria. The filters are then placed, bacteria side up, on plates of the above medium. Once colonies have arisen, usually within 18–24 h at room temperature and longer for lower temperatures, the plates can be examined in a dark room. Luminous colonies can then be picked (sterile wooden toothpicks are suitable for this purpose) and streaked for isolation on fresh plates of the same medium. Use of a red light, such as a photographic darkroom light, on a variable intensity control can make the picking of luminous colonies easier; by adjusting the red light, colonies of nonluminous bacteria can be made to appear reddish, whereas luminous colonies are blue due to their luminescence. Samples collected from warm waters and incubated at room temperature are more likely to yield V. harveyi and related luminous Vibrio species, as well as A. fischeri, P. leiognathi, and P. mandapamensis, whereas cold seawater samples plated and incubated at lower temperatures are likely to yield A. logei, P. kishitanii, P. phosphoreum, and S. hanedai. It should be noted that some strains of A. logei and S. hanedai grow well but do not produce light at room temperature; attempts to isolate these and other psychrotrophic bacteria should be carried out at 15  C. Also, some bacteria, such as A. salmonicida, may not produce visible levels of light unless aldehyde is added to the medium (Fidopiastis et al. 1999); a simple screening approach for finding these bacteria is, once the observer is dark adapted, to add a drop of decyl aldehyde to the underside of the lid of the plates used for plating environmental samples and look for previously dark colonies that then become luminous. Enrichment from fish or squid can be made using fresh samples and sterile seawater or frozen samples with natural, unsterilized seawater (e.g., Ast and Dunlap 2005). The tissue, preferably with the skin on, is placed in a tray, skin up, covered halfway with seawater, incubated, and observed daily in the dark for luminous spots (see > Fig. 13.5), which arise in one to a few days. These spots, colonies of luminous bacteria, can then be picked and streaked for isolation on the medium described above containing 4 % agar. From fish and squid, a variety of different species of luminous bacteria can be isolated, especially when different incubation temperatures, such as 4  C, 15  C, and 22  C, are used. Various crustaceans (e.g., gammarid and caprellid amphipods) are suitable sources for luminous bacteria, as they can

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become infected with luminous (and nonluminous) 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 LSW-70 agar plates. 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.

Cultivation The easily prepared complete medium, LSW-70 (Dunlap et al. 2004), detailed above, is suitable for the growth and luminescence of all culturable luminous bacteria. 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 marine habitats. Previously, a commonly used complete medium was SWC, prepared with natural seawater diluted to 70 % or 75 %, 5 g per liter of tryptone or peptone, 3 g per liter of yeast extract and 3 ml per liter of glycerol, and with 15 g per liter of agar for solid medium. Traditionally, SWC was buffered with 50 mM Tris or HEPES, or 1 g per liter of solid calcium carbonate was incorporated into the agar medium to control acid production (Nealson 1978). Acid production during growth in SWC, which can lead to death of the cells, results, however, from the presence of glycerol, and elimination of this component avoids the problem (Hill 1928; Johnson and Shunk 1936; Dunlap et al. 1995) with no major effect on growth or luminescence. Nealson (1978) listed and compared various formulations for complete and minimal media. Artificial seawater can be prepared according to the formulation of MacLeod, as described by Nealson (1978), or for routine culture work, a commercial aquarium marine salt mix can be used. Procedures for preparing minimal media have been described by Nealson (1978). Growing marine luminous bacteria in liquid medium may require a low agitation rate, since strong aeration can cause some species to clump at low population density.

Storage Luminous bacteria have been revived from sealed glass ampoules after more than 80 years and 28 years of storage (Figge et al. 2011; Haneda 1981). Storage of luminous bacteria on agar slants or in agar stabs is suitable for only short periods, that is, days; longer-term storage on media is not recommended, as dim and dark variants easily arise with most 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. 1984). Perhaps most convenient and effective for retaining strains in their original state is storage at ultralow temperature, for example, 75  C to 80  C, in a cryoprotective medium. An effective cryoprotective medium for the storage of luminous

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bacteria is filter-sterilized double strength deep freeze medium (2X DFM), prepared with 1 % w/v yeast extract, 10 % dimethyl sulfoxide (DMSO), 10 % glycerol, and 02 M K2HPO4/NaH2PO4 (pH 70). 2X DFM, originally developed by R. Rodriquez for storing yeast, was recommended to us several years ago by E. F. DeLong. Storage using 2XDFM works well for all species of luminous bacteria (P. Dunlap, pers. obs.). For storage of a strain, a dense culture is prepared by growing the strain in a complete liquid medium (e.g., LSW-70 broth) with aeration overnight or longer, to attain a dense population, adding 0.5 ml each of the culture and 2X DFM to cryovials, briefly vortexing to mix, allowing the mixture to stand for 5 min, and then placing the vial into the ultralow temperature freezer. Commercial cell freezing containers containing isopropanol, which allow a slow rate of cooling, work particularly well. Quick freezing in an ethanol bath kept in the ultralow temperature freezer or chilled with dry ice works well, but the ethanol can cause the labeling on tubes to smear. Cultures of luminous bacteria stored in this manner retain viability apparently indefinitely when the vials are kept at constant ultralow temperature.

Identification The taxonomy of the marine luminous bacteria and their relationships to other marine enterobacteria were established during the 1970s and 1980s through the use of an array of diagnostic physiological, biochemical, and molecular traits (Reichelt and Baumman 1973; Reichelt et al. 1976; Baumann and Baumann 1981; Baumann and Baumann 1981). Very substantial progress was made through this work in clarifying the genus and species diversity of luminous bacteria, and that work established a foundation for understanding the ecological distributions and evolutionary relationships of these bacteria. At a practical level, the use at the time of as few as 10–25 phenotypic traits allowed the identification of many of the commonly encountered species of marine luminous bacteria (Nealson 1978; Baumann and Baumann 1981; Hastings and Nealson 1981). More recently, however, many of the entities thought to be single species based on these phenetic traits have been found to represent multiple, evolutionarily distinct lineages, that is, separate species and genera, when examined by molecular phylogenetic criteria (e.g., Ast and Dunlap 2005; Ast et al. 2009). Consequently, current methods for the rapid and accurate identification of luminous bacteria and for descriptions of new species increasingly are based on phylogenetic analysis of gene sequences, which is now inexpensive, rapid, and highly accurate. For rapid identifications, sequence analysis of just the luxA or luxB genes often is adequate for good provisional identifications and can be supplemented or replaced by analysis of the sequence of gyrB. A multilocus approach using housekeeping genes such as the 16S rRNA gene, gyrB, pyrH, recA, rpoA, and glnA has proven very effective for robustly separating closely related luminous bacteria and for revealing the rare instances in which a strain apparently has acquired lux genes horizontally.

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Information on primers and amplification procedures for these and other genes can be found in Ast et al. (2009), Thompson et al. (2005), Urbanczyk et al. (2008), Fischer Le Saux et al. (1999), and Peat et al. (2010). Together with housekeeping genes, sequence analysis of the lux genes has proven particularly valuable for new species descriptions and rapid identification of newly isolated strains (e.g., Ast and Dunlap 2004; Haygood 1990; Ast and Dunlap 2005; Thompson et al. 2005; Ast et al. 2007b; Urbanczyk et al. 2007; Ast et al. 2009). Complete characterization of a new species of luminous bacteria, however, should include more than just a multigene phylogenetic analysis. Diagnostic biochemical and morphological traits, DNA hybridization analysis, determination of the mol% G + C ratio, fatty acid profile analysis, and comparative genomic analysis such as amplified fragment length polymorphism (AFLP) or repetitive extragenic polymorphic PCR (repPCR), in the context of the bacterium’s ecology (e.g., Ast et al. 2007a), provide a more complete description suitable for new species. Furthermore, the examination of multiple independent isolates of the new entity and the inclusion in the analysis of the type strains of all closely related and relevant species are critically important for accurate and definitive work.

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Luminous Bacteria Shilo M, Yetinson T (1980) Physiological characteristics underlying the distribution patterns of luminous bacteria in the Mediterranean Sea and the Gulf of Elat. Appl Environ Microbiol 38:577–584 Shimada T, Arakawa E, Itoh K, Kosako Y, Okitsu T, Yamai S, Nishino M, Nakajima T (1995) Causative agent of the so–called ‘‘light disease of shrimps’’ is luminescent Vibrio cholerae non–O1. Nippon Saik Z 50:863–870 Showalter RE, Martin MO, Silverman MR (1990) Cloning and nucleotide sequence of luxR, a regulatory gene controlling bioluminescence in Vibrio harveyi. J Bacteriol 172:2946–2954 Sicard M, Hering S, Schulte R, Gaudriault S, Schulenburg H (2007) The effect of Photorhabdus luminescens (Enterobacteriaceae) on the survival, development, reproduction and behaviour of Caenorhabditis elegans (Nematoda Rhabditidae). Environ Microbiol 9:12–25 Silverman M, Martin M, Engebrecht J (1989) Regulation of luminescence in marine bacteria. In: Hopwood DA, Chater KF (eds) Genetics of bacterial diversity. Academic, London, pp 71–86 Singleton RJ, Skerman TM (1973) A taxonomic study by computer analysis of marine bacteria from New Zealand waters. J R Soc New Zealand 3:129–140 Small ED, Koka P, Lee J (1980) Lumazine protein from the bioluminescent bacterium Photobacterium phosphoreum. Purification and characterization. J Biol Chem 255:8804–8810 Smith SK, Sutton DC, Fuerst JA, Reichelt JL (1991) Evaluation of the genus Listonella and reassignment of Listonella damsela (Love et al.) MacDonell and Colwell to the genus Photobacterium as Photobacterium damsela comb. nov. with an emended description. Int J Syst Bacteriol 41:529–534 Sparks JS, Smith WL, Dunlap PV (2005) Evolution and diversification of a sexually dimorphic luminescent system in ponyfishes (Teleostei: Leiognathidae), including diagnoses for two new genera. Cladistics 21:305–327 Spencer R (1961) Chitinoclastic activity of the luminous bacteria. Nature 190:938 Stacy AR, Diggle SP, Whiteley M (2012) Rules of engagement defining bacterial communication. Curr Opin Microbiol 15(2):155–162 Stevens AM, Greenberg EP (1997) Quorum sensing in Vibrio fischeri: essential elements for activation of the luminescence genes. J Bacteriol 179:557–562 Sun W, Cao JG, Teng K, Meighen EA (1994) Biosynthesis of poly–3– hydroxybutyrate in the luminescent bacterium, Vibrio harveyi, and regulation by the lux autoinducer, N–(3–hydroxybutanoyl) homoserine lactone. J Biol Chem 269:20785–20790 Sung ND, Lee CY (2004) Coregulation of lux genes and riboflavin genes in bioluminescent bacteria of Photobacterium phosphoreum. J Microbiol 42:194–199 Surete MG, Miller MB, Bassler BL (1999) Quorum sensing in Escherichia coli, Salmonella typhimurium, and Vibrio harveyi: a new family of genes responsible for autoinducer production. Proc Natl Acad Sci USA 96:1639–1644 Suwanto A, Yuhana M, Herawaty E, Angka SL (1998) Genetic diversity of luminous Vibrio isolated from shrimp larvae. In: Flegel TW (ed) Advances in shrimp biotechnology. National Center for Genetic Engineering and Biotechnology, Bangkok, pp 217–224 Swartzman E, Kapoor S, Graham AF, Meighen EA (1990a) A new Vibrio fischeri lux gene precedes a bidirectional termination site for the lux operon. J Bacteriol 172:6797–6802 Swartzman E, Miyamoto C, Graham A, Meighen E (1990b) Delineation of the transcriptional boundaries of the lux operon of Vibrio harveyi demonstrates the presence of two new lux genes. J Biol Chem 265:3513–3517 Swartzman E, Silverman M, Meighen EA (1992) The luxR gene product of Vibrio harveyi is a transcriptional activator of the lux promoter. J Bacteriol 174:7490–7493 Swift S, Williams P, Stewart GSAB (1999) N–Acylhomoserine lactones and quorum sensing in proteobacteria. In: Dunny GM, Winans SC (eds) Cell–cell signaling in bacteria. American Society for Microbiology Press, Washington, pp 291–313 Tailliez P, Laroui C, Ginibre N, Paule A, Page`s S, Boemare N (2010) Phylogeny of Photorhabdus and Xenorhabdus based on universally conserved protein– coding sequences and implications for the taxonomy of these two genera. Proposal of new taxa X. vietnamensis sp. nov., P. luminescens subsp.

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14 Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria Dimitry Y. Sorokin1,2 . Horia Banciu3 . Lesley A. Robertson2 . J. Gijs Kuenen2 . M. S. Muntyan4 . Gerard Muyzer2,5 1 Winogradsky Institute of Microbiology, Russian Academy of Sciences, Moscow, Russia 2 Department of Biotechnology, Delft University of Technology, Delft, The Netherlands 3 Faculty of Biology and Geology, Babes-Bolyai University, Cluj-Napoca, Romania 4 Belozersky Institute of Physico-Chemical Biology, Moscow State University, Moscow, Russia 5 Department of Aquatic Microbiology, Institute for Biodiversity and Ecosystem Dynamics, University of Amsterdam, Amsterdam, The Netherlands

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 530 Haloalkaliphilic SOB in Soda Lakes . . . . . . . . . . . . . . . . . . . . . . . 530 Soda Lakes as a Unique Habitat . . . . . . . . . . . . . . . . . . . . . . . . . 530 Enrichment, Isolation, and Cultivation of Haloalkaliphilic SOB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 532 Distribution and Diversity of Haloalkaliphilic Chemolithoautotrophic SOB . . . . . . . . . . . . . . . . . . . . . . . . . . . . 533 Ecophysiology of Aerobic Haloalkaliphilic SOB . . . . . . . . 535 Sulfur Oxidation Mechanisms in Haloalkaliphilic SOB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 540 Specific Physiological Subgroups of the Soda Lake SOB . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 540 Denitrification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 540 Thiocyanate Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 542 Facultatively Alkaliphilic SOB from Hypersaline Alkaline Lakes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 543 Culture-Independent Detection of Haloalkaliphilic SOB in Soda Lake Sediments . . . . . . . . . . . . . . . . . . . . . . . . . . . . 543 Haloalkaliphilic SOB in ‘‘Artificial Soda Lakes’’ . . . . . . . . . 543 Neutrophilic Halophilic SOB from Hypersaline Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544 Chloride-Sulfate Hypersaline Habitats with Neutral pH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544 Enrichment of Halophilic SOB . . . . . . . . . . . . . . . . . . . . . . . . . . 544 Moderately Halophilic Aerobic SOB . . . . . . . . . . . . . . . . . . . . 545 Moderately Halophilic Thiodenitrifying SOB . . . . . . . . . . . 545 Extremely Halophilic Aerobic SOB . . . . . . . . . . . . . . . . . . . . . 545 Extremely Halophilic Denitrifying SOB . . . . . . . . . . . . . . . . . 546 Halophilic Thiocyanate-Oxidizing SOB . . . . . . . . . . . . . . . . . 548 Culture-Independent Analysis of Halophilic SOB in a Hypersaline Lake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 549 Conclusions and Future Prospects . . . . . . . . . . . . . . . . . . . . . . . . . 551

Abstract Chemotrophic sulfur-oxidizing bacteria (SOB) represent an important functional group of microorganisms responsible for the dark oxidation of reduced sulfur compounds generated by sulfidogens. Until recently, only a single genus of halophilic SOB (Halothiobacillus) has been described, and nothing was known about the ability of this group to grow at high pH. Investigation of soda lakes, unique extremely alkaline and saline habitats, led to the discovery of a novel ecotype of natronophilic SOB. In contrast to the previously known neutrophilic ecotype, this group cannot grow at neutral pH, but grows optimally in soda brines at pH values around 10. They were the first chemolithoautotrophs among the described alkaliphiles. The group, so far, includes four novel genera within the Gammaproteobacteria. The genera Thioalkalimicrobium and Thioalkalispira represent low salt-tolerant alkaliphiles tolerating up to 1.5 M Na+. The genus Thioalkalibacter is a high salttolerant facultative alkaliphile. The genus Thioalkalivibrio is the most diverse and includes aerobic extremely salt-tolerant members and moderately salt-tolerant thiocyanate-utilizing and facultatively anaerobic denitrifying strains. The genome sequence of two Thioalkalivibrio strains revealed the presence of a truncated Sox system lacking the SoxCD component which is typical for gammaproteobacterial SOB. Bioenergetic studies of high salt-tolerant Thioalkalivibrio strains showed an obligate sodium dependence for respiratory activity implying the presence of sodium-dependent elements. Investigation of hypersaline inland chloride-sulfate lakes and hypersaline brines of marine origin with neutral pH revealed an unexpectedly high culturable diversity of halophilic obligately chemolithoautotrophic SOB comprising seven different groups within the Gammaproteobacteria. Two groups of strictly aerobic moderate halophiles were represented by the known genera Halothiobacillus and Thiomicrospira. Under denitrifying conditions and with thiocyanate as e-donor, three novel groups of

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Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

moderately halophilic SOB were represented by the genera Thiohalomonas, Thiohalophilus, and Thiohalobacter. At 4 M NaCl, two groups of extremely halophilic SOB (a type not known before among the SOB) had been discovered. The obligately aerobic extreme halophiles comprised a novel genus Thiohalospira, and the facultatively anaerobic nitrate-reducing extreme halophiles—a novel deep-lineage genus Thiohalorhabdus. Overall, the investigation of hypersaline and (halo)alkaline habitats uncovered a novel and diverse world of extremophilic SOB with properties previously unknown for chemolithoautotrophic bacteria.

Introduction Chemolithotrophic SOB play an important role in the cycling of different chemical elements in natural and man-made environments because of their capacity to transform various sulfur and nitrogen compounds and their contribution to secondary production of organic matter. They are widely distributed in various habitats, associating primarily with sulfide-oxygen interface layers, where they compete with chemical of sulfide oxidation by oxygen. The reaction of complete oxidation of sulfide or thiosulfate to sulfate (eight electrons) is among the most attractive for chemosynthesis, and it is therefore not surprising that chemolithotrophic SOB can be found in many different groups of the Prokaryotes. Currently, lithoautotrophic SOB are mostly found among the Proteobacteria (alpha, beta, gamma, and epsilon subdivision; see > Chap. 15, ‘‘Colorless Sulfur Bacteria’’ in this volume; Robertson and Kuenen 2006; Kelly and Wood 2000). Examples outside the Proteobacteria are represented by the Grampositive bacteria of the genus Sulfobacillus (Johnson et al. 2008), thermoacidophilic Crenarchaeota (Kletzin et al. 2004), and deep bacterial lineages, such as Aquificales and Thermus (Huber and Eder 2006; da Costa et al. 2006) (> Table 14.1). According to their response to pH, the known sulfur-oxidizing species include acidophiles (optimum pH Fig. 14.1. Our studies of alkaliphilic SOB included Siberian, Kenyan, Mongolian, North American soda lakes and hypersaline alkaline lakes in Egypt. The main characteristics of the lakes studied are given in > Table 14.2. The unusually high pH establishes several specific chemical conditions important for microbial sulfur cycling. Sulfide is present entirely in the ionic form as HS , which, in contrast to H2S, cannot freely cross the membrane, and therefore is not as toxic as H2S. Accordingly, alkaliphilic SOB can tolerate much higher substrate concentrations than their neutrophilic counterparts. Next, sulfide can react chemically with insoluble sulfur forming soluble polysulfides that are chemically stable at high pH. At aerobic conditions, polysulfide is rapidly oxidized to thiosulfate, while at anaerobic conditions, it is stable and hence can accumulate to high concentrations representing a true substrate for sulfur-reducing prokaryotes. High alkalinity is favorable for SOB because of a buffering effect against the

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

14

. Table 14.1 Lithotrophic SOB diversity Taxa

Representative species

Obligate/facultative

Anaerobic growth with NOx

pH/temp/salt

Paracoccus

P. denitrificans

F

+

n/m/nh

Acidiphilium

A. acidiphilum

F

–

ac/m/nh

Starkeya

S. novella

F

–

n/m/nh

Thioclava

Tcv. pacifica

F

–

n/m/mr

Thiobacillus

Tb. thioparus

O

–

n/m/nh

Thiobacter

Tbc. subterraneus

O

–

n/m/nh

Thiomonas

Ts. intermedius

F

–

ac/m/nh

Magnetospirillum

M. magnetotacticum

F

–

n/m/nh

Thermothiobacillus

Tt. tepidarius

O

–

n/t/nh

Acidithiobacillus

Atb. thiooxidans

O

–

ac/m/nh

Halothiobacillus

Htb. halophilus

O

–

n/m/h

Thiomicrospira

Tm. crunogena

O

–

n/m/mr–h

Thiovirga

Thv. sulfuroxydans

O

–

n/m/nh

Thioalkalivibrio

Tav. versutus

O

–

al/m/h

Thioalkalimicrobium

Tam. aerophilum

O

–

al/m/h

Thioalkalispira

Tas. microaerophila

O

–

al/m/h

Thiohalobacter

Thb. halophilus

O

–

Thiohalophilus

Thp. thiocyanoxidans

O

+

n/m/h

Thiohalomonas

Thm. denitrificans

O

+

n/m/h

Thiohalospira

Ths. sibirica

O

–

n/m/eh

Thiohalorhabdus

Thr. denitrificans

O

+

n/m/eh

‘‘Thiobacillus prosperus’’

‘‘Thiobacillus prosperus’’ O

–

ac/m/nh

Thiothrix

Thiothrix ramosa

F

–

n/m/nh

Beggiatoa

B. alba

F

–

n/m/nh

Alkalilimnicola

A. ehrlichii

F

+

al/m/h

Sulfurimonas

Sm. denitrificans

O

+

n/m/mr

Sulfurovum

Sr. lithotrophicum

O

–

n/m/mr

Sulfuricurvum

Sc. kujiense

O

Genus

Proteobacteria Alpha

Beta

Gamma

Epsilon

n/m/mr

Arcobacter

Cand ‘‘Arc. sulfidicus’’

O

–

n/m/mr

Firmicuta

Sulfobacillus

Sb. thermosulfidooxidans F

–

ac/t/nh

Deep-lineage Bacteria

Hydrogenivirga

Hv. caldilitoris

O

–

ac/t/nh

Hydrogenobacter

Hgb. thermophilus

F

–

ac/t/nh

Sulfurihydrogenibium

Sh. azorense

O

–

ac/t/nh

Thermocrinis

Tc. ruber

F

–

ac/t/nh

Aquifex

Aq. pyrophilus

O

–

ac/t/nh

Sulfolobus

Sl. acidocaldarius

F

–

ac/t/nh

Acidianus

Ac. ambivalence

F

–

ac/t/nh

Sulfurococcus

S. yellowstonensis

F

–

ac/t/nh

Stygiolobus

St. azoricus

F

–

ac/t/nh

Crenarchaeota

F fac. autotrophic, O obl. autotrophic, ac acidophilic, n neutrophilic, al alkaliphilic, m mesophilic, t thermophilic, nh freshwater, mr marine, h moderately halophilic, eh extremely halophilic

531

532

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Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

. Fig. 14.1 Typical small shallow soda lake (Kulunda steppe, Altai, Russia)

. Table 14.2 Soda lakes studied pH

Total carbonate alkalinity (M)

20–220

9.5–11.0

0.12–1.16

200–380

9.5–10.3

0.11–0.75

Mono Lake, Searles Lake, Owens Lake

90–220

9.7

0.5–1.0

Hilganta, Hadyn, Ulan-Nor, Gorbunka

5–40

9.5–10.2

0.02–0.11

20–380

9.3–10.6

0.02–5.2

9.2–10.5

0.02–1.20

Area

Examples

Total salt, g l

Kenya

Magadi, Bogoria, Crater Lake, Elmenteita

Wadi Natrun (Egypt)

Rozita, Beida, Gaar, Fazda, Zugm, Hamra

California Transbaikal Steppe (Southeastern Siberia, Russia) Kulunda Steppe (Altai region, Russia)

Cock soda lake Salt Lake Steppe system Tanatar lake system Bitter lake system

Northeastern Mongolia

Hotontyn, Shar-Burdiin, Dzun-Uldziit, Baga-Nur 5–360

sulfuric acid produced during sulfur oxidation in the periplasm. On the other hand, domination of carbonate over bicarbonate at pH values close to 10 and above is unfavorable for CO2/HCO3 assimilation, setting a limitation on the growth of autotrophic alkaliphiles at extremely high pH. Recent microbiological analysis of the soda lakes by both traditional and culture-independent molecular techniques revealed the remarkable fact that, despite the double extreme conditions of salt and pH, fully structured and active microbial communities are present, even in saturated alkaline brines (Imhoff et al. 1979; Grant and Tindall 1986; Jones et al. 1998; Zavarzin et al. 1999; Zavarzin and Zhilina 2000; Humayoun et al. 2003; Sorokin et al. 2004a; Ma et al. 2004; Rees et al. 2004; Mesbah et al. 2007; Zavarzin 2007; Foti et al. 2007, 2008). Among them, the main functional groups of anaerobes, such as fermentative, acetogenic, methanogenic, and sulfate-reducing bacteria, represented by unique haloalkaliphilic species, have recently been isolated and identified. The microbial sulfur cycle seems to be one of the most active in the soda lakes, with anaerobic phototrophic purple sulfur bacteria, chemolithotrophic SOB, and respiratory sulfidogenic

1

alkaliphiles as the main participants (Isachenko 1951; Imhoff et al. 1979; Zhilina et al. 1997; Gorlenko et al. 1999; Sorokin et al. 2001a, 2010).

Enrichment, Isolation, and Cultivation of Haloalkaliphilic SOB Following the published examples (Tindall 1988; Horikoshi 1991), our first attempts to enrich alkaliphilic SOB from soda lake sediments were made using a culture medium containing 20 mM thiosulfate as energy source, 10 g l–1 of sodium carbonate, 0–50 g l–1 NaCl, 0.5 g l–1 NH4Cl, and 1 g l–1 of K2HPO4. Sodium carbonate was added after sterilization, and the pH was adjusted to 10 by HCl. Some growth was observed on this medium with an inoculum from the Hadyn soda lake in Tuva (> Table 14.2), but the resulting cultures were not stable, and, in particular, a drop in pH below 9 was followed by heavy sulfur precipitation. An optimization resulted in the formulation of a mineral medium, containing 0.6–4.0 M total Na+ based on carbonate buffer. This mineral base can be sterilized in closed

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

bottles and has a stable pH 10. For the hypersaline alkaline lakes in which NaCl was the dominant salt (e.g., the Wadi Natrun lakes, Stamp Lake, Searles Lake), the amount of sodium carbonate in the base 4 M Na+ medium was reduced by substituting NaCl with the ratio of sodium carbonates to NaCl 1 M:3 M—2 M:2 M. This medium provided sufficient alkalinity so that even when a dense sulfur-oxidizing culture producing high amounts of sulfuric acid from thiosulfate developed, the pH remained above 9.5. Potassium nitrate was provided as the source of nitrogen and potassium. In addition, in some cases, ammonium chloride was added after sterilization at low concentration (2 mM) to ensure enrichment of species unable to utilize nitrate. The low concentration is dictated by the almost complete deprotonation of ammonium to highly toxic, free ammonia at pH 10. After sterilization, the alkaline mineral base medium was supplied with 20–80 mM of thiosulfate, 1 ml per liter of trace elements solution (Pfennig and Lippert 1966), and 1 mM MgCl2·6H2O. The latter formed a soluble basic magnesium carbonate complex [Mg2(OH)2CO3]. The use of this mineral medium allowed successful enrichment and isolation of multiple strains of obligately alkaliphilic SOB with varying degrees of salt tolerance. The preparation of solid alkaline media is complicated by the chemical instability of agar at high pH at temperatures above 50  C and by the low solubility of sodium carbonate. Therefore, the preparation of solid medium involves mixing equal volumes of sterile thoroughly prewashed 4 % (w/v) agar and mineral base medium in double strength at 50  C. Obviously, the maximum soda concentration in the final solid medium was then limited to 2 M Na+. The isolation strategy was based on the following procedure. Positive enrichment cultures were subcultured several times with 1:100 inocula to obtain a stable active culture. This stable culture was serially diluted, and the successive dilutions were plated. Sometimes, especially when the low-salt medium (0.6 M Na+) was used, it was necessary to make serial dilutions immediately without preliminary enrichment because of the high grazing activity of protozoa. This approach was also useful in combining enrichment, enumeration, and isolation in a single procedure. Moreover, it later appeared that the efficiency of colony formation of haloalkaliphilic SOB was low in comparison with their growth on liquid media. Therefore, in most cases, the dilution-to-extinction approach was used to isolate the dominant species growing in liquid cultures. Growth was monitored by measuring thiosulfate consumption (iodimetric titration in presence of acetic acid) and by microscopical observation of cell growth. Some variations in the selective enrichment strategy, such as the use of sulfide and thiocyanate instead of thiosulfate as substrate or nitrogen oxides instead of oxygen as electron acceptors, will be discussed below.

Distribution and Diversity of Haloalkaliphilic Chemolithoautotrophic SOB Enrichment on low-salt alkaline medium under fully aerobic conditions using inocula from the Siberian low-salt soda lakes

14

gave the first indication of the presence of aerobic SOB capable of stable growth at pH 10 (Sorokin et al. 2000). Two pure cultures, strains AL2 and AL3, isolated from lake Hadyn in Tuva (Russia) became the reference type strains for two different groups of the haloalkaliphilic SOB mainly distributed in soda lakes. Subsequent investigation of samples from different geographic locations (> Table 14.2) resulted in the isolation of more than 100 strains of obligately alkaliphilic chemolithoautotrophic SOB. Positive enrichment cultures were only obtained from lakes with pH >9 and not from the salt lakes with pH between 7 and 8.5. They were present in the soda lake sediments at relatively high numbers from 103 to up to 108 cells/g. The cultivation and maintenance of the haloalkaliphilic SOB is not complicated by acid production because of the extremely high buffering capacity of the soda-based medium. For example, full oxidation of 80 mM thiosulfate (20 g/l) resulted in a pH drop from 10 to 9.2 even at the lowest buffering capacity used (0.6 M total Na+). The dense liquid cultures utilizing between 40 and 80 mM thiosulfate remained viable during storage at 4  C for 3–6 months. Most of the strains also survived freeze-drying and deep-freezing storage in 10 % (v/v) glycerol. The soda lake SOB isolates formed four groups within the Gammaproteobacteria described as new genera Thioalkalimicrobium, Thioalkalivibrio (Sorokin et al. 2001a, 2002a, b, c, 2003a, 2004a; Banciu et al. 2004a), Thioalkalispira (Sorokin et al. 2001c), and Thioalkalibacter (Banciu et al. 2008) (> Fig. 14.2). The genus Thioalkalimicrobium with a typical cell morphology (> Fig. 14.3) is closely related to the marine SOB of the Thiomicrospira pelophila cluster. The genus Thioalkalivibrio (> Fig. 14.4) is a member of the family Ectothiorhodospiraceae, which includes mainly halophilic and haloalkaliphilic species (Imhoff 2006). The genus Thioalkalispira has recently been classified into a separate family Thioalkalispiraceae (Mori et al. 2011), although phylogenetically it is also related to the Ectothiorhodospiraceae. The genus Thioalkalibacter is forming a separate lineage within the group of Halothiobacillus and its appearance made it quite clear that the latter needs a reclassification into a family including three separate genera: (a) the low salt-tolerant marine species Htb. neapolitanus and Htb. kellyi, (b) the truly halophilic species Htb. halophilus and Htb. hydrothermales, and the facultatively alkaliphilic halophile Thioalkalibacter halophilus. The genera Thioalkalimicrobium, Thioalkalispira, and Thioalkalivibrio accommodate exclusively obligately alkaliphilic members, with the exception of the facultatively alkaliphilic Thioalkalivibrio halophilus. Thialkalibacter halophilus is a facultatively alkaliphilic halophile. In general, the Thioalkalimicrobium group dominated enrichment cultures from lowsaline Asian soda lakes. It also could be enriched from the hypersaline soda lakes, but only from fresh samples and in low-salt culture medium. The genus Thioalkalivibrio dominated enrichments from hypersaline soda lakes and from aged samples and was always dominant when the enrichment medium contained >1.5 M total Na+. Therefore, the most important selective force influencing the composition and survival of alkaliphilic SOB in soda lakes appeared to be the salt

533

534

14

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria Thioalkalivibrio ALR9, EU709854

0.05

Thioalkalivibrio ALMg2, AY079149 Thioalkalivibrio versutus AL2T, AF126546 Thioalkalivibrio ALMg14, EU709869 Thioalkalivibrio jannaschii ALM2T, AF329083 Thioalkalivibrio AKL15, EU709871 Thioalkalivibrio ALR16, EU709875 Thioalkalivibrio ALJ6, GU735092 Mono Lake, clone 623J-18, AF507820 Thioalkalivibrio ALOw1, GQ863491 100

99

Hypersaline mat, clone E2aA03, DQ103664 Thioalkalivibrio thiocyanoxidans ARh2T, AF302081 Thioalkalivibrio nitratis ALJ12T, AF126547 Thioalkalivibrio AKL11, EU709870 Thioalkalivibrio ALR4, EU709850

Thioalkalivibrio K90mix, EU709865 Thioalkalivibrio ASL10, DQ900624 Thioalkalivibrio ALJ22 Thioalkalivibrio ALJ15, GU735093 Thioalkalivibrio ALE10, GU735088 Thioalkalivibrio ALSr1, GU735087 81 Thioalkalivibrio ALJ24, EU709867 76 Thialkalivibrio halophilus HL17T, AY346464 87 Thioalkalivibrio ALR6, EU709852 85 Thioalkalivibrio ALR20, EU709876 Thioalkalivibrio ALE20, EU709852 Thioalkalivibrio AKLD2 95 Thioalkalivibrio paradoxus ARh 1T, AF151432 95 Thioalkalivibrio nitratireducens ALEN 2T, AY079010 Wadi Natrun, clone HWB-90, DQ432366 76 96 Mono Lake, clone 635J-54, AF507822 Thioalkalivibrio ALN1, EU709866 Thioalkalivibrio denitrificans ALJDT, AF126545 100 Thioalkalivibrio thiocyanodenitrificans ARhDT, AY360060 81 Thioalkalivibrio “sulfidophilus” HL-EbGr7T, EU709878 100 Thioalkalivibrio “sulfidophilus” ALJ 17, GU735094 Ectothiorhodospira mobilis DSM 237T, X93481 100 Ectothiorhodospira halalkaliphila BN 9903T, X93479 Halorhodospira halophila SL1T, M26630 100 89 Halorhodospira abdelmalekii DSM 2110T, X93477 Marine cold seep, clone JT58-36, AB189351 Thioalkalispira microaerophila ALEN 1T, AF481118 81 Thioalkalibacter ALOw3 99 Thioalkalibacter ALOw2, GQ863492 Thioalkalibacter halophilus ALCO1T, EU124668 90 94 Halothiobacillus halophilus DSM 6132T, U58020 Halothiobacillus neapolitanus DSM 15147T, AF173169 89 Thiovirga sulfuroxydans SO07, AB118236 Thiomicrospira crunogena ATCC 35932T, L40810 93 100 Thiomicrospira kuenenii JB-A1T, AF013978 100 Thioalkalimicrobium microaerophilum ASL8-2T, DQ900623 Thiomicrospira pelophila DSM 1534T, L40809 100 Thiomicrospira thyasirae DSM 5322T, AF016046 100 Thioalkalimicrobium sibiricum AL7T, AF126549 83 Thioalkalimicrobium cyclicum ALM1T, AF329082 Thioalkalimicrobium aerophilum AL3T, AF126548 100 Thioalkalimicrobium ALE5, GU735086 100 Thioalkalimicrobium ALE3, GU735085 89

100

. Fig. 14.2 Phylogenetic position of haloalkaliphilic SOB from soda lakes (in bold) within the Gammaproteobacteria based on 16S rRNA sequence analysis. The strain abbreviations are as follows: AL, strains from Transbaikal region (Russia), AKL, strains from Kulunda Steppe (Altai, Russia), ALMg strains from n-e Mongolia, ALJ strains from Kenya, ALE strains from Wadi Natrun in Egypt, ASL strain from Soap Lake (USA), ASLr strain from Searles Lake (USA), ALOw strains from Owens lake (California), ALN1 extremely haloalkaliphilic nitrate-reducing strain from Wadi Natrun, ALR strains from a lab–scale sulfide-oxidizing bioreactor, HL-EbGr7 a strain from a full-scale sulfide-oxidizing bioreactor. In red are the strains with sequenced genome and in pink are strains which genome sequencing is in progress. Numbers at the nodes indicate the percentage of bootstrap values for the clade of this group in 1,000 replications (the values for maximum-likelihood method are given in parentheses). Only values above 70 % are shown. Bar, 5 substitutions per 100 nucleotides (nt)

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

14

. Fig. 14.3 Cell morphology of the genus Thioalkalimicrobium. (a, c–e) total electron microphotographs; (b, f) thin sections; (a–b) Tm. aerophilum; (c) Tm. sibiricum; (d) Tm. microaerophilum (e–f) Tm. cyclicum, Cs caboxysomes, N nucleoide. Bars: 0.5 mm

concentration (> Table 14.3). The genus Thioalkalivibrio is apparently the most widely distributed SOB type in soda lakes. Among its multiple isolates, several large geographical populations each containing about 20 isolates could be distinguished (Foti et al. 2006). Extremely salt-tolerant isolates from the Kulunda Steppe and Mongolia are clustered together and also related to the strains from Mono Lake and Soap Lake in northern America, while most of the Wadi Natrun isolates are clustering with Tv. halophilus. Such clustering correlates with the anionic composition of the brines (see below). The only other culturable evidences on haloalkaliphilic SOB came from the dry soda lake Texcoco in Mexico (Granada et al. 2009), although none of the isolates were characterized. The presence of the SOB related to the genus Thioalkalivibrio detected in 16 S rRNA gene-based libraries has been reported

over the past 10 years from various saline habitats, including marine sources, hypersaline mats (Sørensen et al. 2005), and soda lakes (Baumgarte 2003; Humayoun et al. 2003; Rees et al. 2004; Mesbah et al. 2007). Marine clones, however, had only 90–95 % sequence similarity to the species of Thioalkalivibrio and therefore might belong to another genus.

Ecophysiology of Aerobic Haloalkaliphilic SOB The main properties of the haloalkaliphilic SOB are presented in > Tables 14.4 and > 14.5. In their basic physiology, the soda lake SOB are typical chemolithoautotrophs using the Calvin cycle for inorganic carbon fixation (Tourova et al. 2006, 2007) and growing best with thiosulfate in batch cultures and sulfide/polysulfide

535

14

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

a

b

10 µm

S

c

10 µm

d

e

f

N Pg g

1 µm

536

h

N Cs . Fig. 14.4 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

in substrate-limited chemostats. Their main difference with the neutrophilic SOB is the ability to grow chemolithoautotrophically at combined high pH and high salinity. The potential for this kind of metabolism was not known before. So, the main characteristic of the soda lake SOB is their preference for a sodium carbonate–based environment with optimal growth at a pH around 10. The pH-controlled continuous cultures (Sorokin et al. 2003a; Banciu et al. 2004a) demonstrated growth

pH maximum of 10.5 for both Thioalkalimicrobium and Thioalkalivibrio type species, while cells remained metabolically active up to pH 11 (> Fig. 14.5). A possible explanation is that their growth, that is, Ci assimilation, was limited by inorganic carbon availability because of the domination of unavailable carbonate over available bicarbonate, as has been suggested for alkaliphilic cyanobacteria (Kaplan et al. 1982), whereas energy metabolism can still proceed at higher pH values.

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

14

. Table 14.3 Results of enrichment and isolation of two different types of haloalkaliphilic SOB from soda lakes (pH 10) Low-salt medium (0.6 M Na+)

High-salt medium (2–4 M Na+)

Lake

MPN

Tm

Tv

MPN

Tm

Tv

Lake Hadyn (Tuva)

nd

nd

nd

1

1

nd

Kunkur Steppe (Siberia)

6

10

14

4

nd

nd

nd

Northeast Mongolia

106

20

0

105

0

20

Lake Borzinskoe (Siberia)

106

0

2

107

0

1

Kulunda Steppe (Siberia)

8

10

3

6

8

10

0

7

Kenya (Rift Valley)

106

3

20

106

0

5

Egypt (Wadi Natrun)

106

4

5

106

0

23

Mono Lake (California)

nd

1

0

nd

0

1

3

Abbreviations: MPN maximum cell number/cm of sediment, Tm number of isolated Thioalkalimicrobium strains, Tv number of isolated Thioalkalivibrio strains, and nd not determined

. Table 14.4 Basic properties of haloalkaliphilic SOB from soda lakes Property

Thioalkalimicrobium spp.

Thioalkalispira microaerophila

Thioalkalivibrio spp.

Thioalkalibacter halophilus

Number of species

4

1

9

1

Number of isolates

43

1

95

3

Close relative

Thiomicrospira pelophila

No

Ectothiorhodospira

Halothiobacillus

Cell morphology

Rods and spirilla

Spirilla

Rods, vibrios, spirilla and cocci

Rods

Sulfur compounds oxidized HS , Sn2 , S2O32

HS , Sn2 , S2O32

HS , Sn2 , S2O32 , S8, SO32 , S4O62 , SCN

HS , Sn2 , S2O32

Electron acceptors

O2

Microaerophile

O2, NO3 , NO2 , N2O

O2

pH optimum

9.5–10.0

10.0

10.0–10.2

8.5

Upper salt limit

1.5 M Na+

1.4 M Na+

4.3 M Na+

3.5 M Na+

Maximal specific growth rate

0.33 h

Maximal growth yield with S2O32

3.5 mg protein mmol 1

5.8 mg protein mmol

Rate of HS oxidation

Extremely high

Low

Low

High

Dominant compatible solute

Ectoine

nd

Glycine betaine

Ectoine

Yellow membrane pigment –

+

+

–

RuBisCO

Form Ia

Form Ia

Form Ia

1

Form Ia

0.08 h

1

0.25 h 1

1

6.5 mg protein mmol

0.22 h 1

1

3.5 mg protein mmol 1

nifH

–

+

–

–

Distribution

Asia, Africa, North America

Egypt

Asia, Africa, North America

Asia, North America

Apart from extreme pH, the salt concentration and ion composition of the brines are very important for the soda lake SOB growth and activity. The genera Thioalkalimicrobium and Thioalkalispira include only moderately salt-tolerant species, Thioalkalivibrio includes both moderately and high salt-tolerant

representatives, and Thioalkalibacter halophilus is a high salttolerant organism. Furthermore, the main characteristic of the soda lake microbes and SOB, in particular, is their preference of sodium carbonates over NaCl. In this respect, they must not be called ‘‘haloalkaliphiles’’ (the traditional term) but rather

537

14

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

. Table 14.5 Respiratory activity in haloalkaliphilic SOB grown with thiosulfate or thiocyanate at pH 10 Thioalkalimicrobium Substrate 2

Thiosulfate (S2O3 )

Thioalkalivibrio

Thioalkabacter halophilus

pH opt

V

N

pH opt

V

N

9–10

2.5–5.2

40

9–10

0.15–1.10

60

pH opt

V

Sulfide (HS )

9–10

2.3–5.2

40

9–10

0.15–1.50

60

10

1.20

Polysulfide (S8 2)

10

1.1–3.0

38

10

0.20–0.90

55

10

1.60

Elemental sulfur (S8)

10

0–0.2

40

10–10.5

0.08–0.60

60

8.0

0.11

Sulfite (SO3 )

0

28

10

0–0.20

40

10.0

Trithionate (S3O6 2)

0

9

9

0–0.20

20

0

0–1.1

40

9

0.05–0.5

60

0

0

9

9

0.1–0.80

20

10

0.09–0.40

9

10

0.09

1

2

Tetrathionate (S4O6 2)

9

2

Pentathionate (S5O6 ) Thiocyanate (SCN )

No growth and respiration

Carbon disulfide (CS2)

0.16

0 No growth and respiration

Abbreviations: V respiration rate, mmol of O2 (mg of protein min) 1; N number of tested strains

b

µ

3

0.2 0.15

2.5

0.1 2

0.05 0 8

9

Y (g prot/Mole S2O32-)

3.5 Y

7

0.25

6 5

0.2

4 0.15 3 0.1 2 0.05

1

0

1.5 11

10

7

8

9 pH

pH

c

µ

Y

0.3 0.25

Thioalkalivibrio versutus AL 2

4

10

y (g prot/Mole S2O32-)

Thioalkalimicrobium aerophilum AL 3 0.35

m (h-1)

a

m (h-1)

0 11

4 Tm

Activity (µmol/(mg prot min)

538

3

2

Tv

1

0 6.5

7.5

8.5

9.5

10.5

11.5

pH

. Fig. 14.5 pH profiles for growth rate (m) and growth yield (Y) of alkaliphilic SOB measured in pH-controlled thiosulfate-limited continuous culture (0.6 M total Na+) (a, b) and for the thiosulfate-dependent respiration of washed cells (c) in Thio- alkalimicrobium and Thioalkalivibrio

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

‘‘natronophiles’’ (soda-loving organisms). The isolation of two truly haloalkaliphilic species, Tv. halophilus and Thioalkalibacter halophilus, which differed from all the other soda lake SOB by their high demand for NaCl and their facultative alkaliphily, showed a very interesting difference in their cellular composition after growth in NaCl-dominated and sodium carbonate–dominated medium at pH 10. The specific concentration of compatible solutes in both organisms was roughly two times lower in the soda-grown cells than in NaCl-grown cells, despite the fact that both media contained the same total Na+ concentration (Banciu et al. 2004a, 2008). This exactly correlated with the difference in osmotic pressure of the two media, which can be easily explained by the electrolytic properties of the two salts (NaCl—strong electrolyte, sodium carbonates—weak electrolytes). So, life at extremely high pH demands extra energy to maintain an unfavorable pH gradient, but the costs of compatible solute synthesis in natronophiles are much lower than in the neutrophilic halophiles. Such a tendency can also be seen in comparison of the growth dynamics of a typical obligate natronophilic Thioalkalivibrio strains from hypersaline soda lakes in Siberia and haloalkaliphilic Thioalkalivibrio isolates from hypersaline alkaline lakes in Wadi Natrun. The former definitely preferred to grow in weak electrolytes and was inhibited by NaCl above 2 M, while the latter grew optimally in presence of at least 2 M NaCl (> Fig. 14.6). Moreover, some of the natronophilic Thioalkalivibrio have another extraordinary difference in comparison to the halophiles. They can still grow well at pH 10 when 90 % of the sodium carbonate was replaced by potassium carbonate (i.e., 3.6 M K+/0.4 M Na+ as carbonates), while halophilic SOB from chloride-sulfate lakes (see below) cannot even grow at a 50 % replacement of sodium for potassium. We also have evidence of the specific requirement for sodium cations for sulfide- and TMPD-dependent respiratory

a

14

activity of the isolated membrane vesicles from Thioalkalivibrio strains pointing to the involvement of sodium-based components in the bioenergetic machinery of natronophilic chemolithoautotrophs (Grischuk et al. 2003). The genus Thioalkalimicrobium includes obligately aerobic, low salt-tolerant, fast-growing, and low-yield strains with extremely high sulfide- and thiosulfate-oxidizing activity. In contrast, the genus Thioalkalivibrio is more phylogenetically and physiologically diverse and accommodates relatively slow growing organisms with a more efficient substrate conversion resulting in higher specific growth yield. They are, in general, more salt-tolerant, with many strains able to grow in saturated soda brines (the core subgroup clustering around the type strain Thioalkalivibrio sp. AL2). Only such strains were selected at a salinity of 2–4 M Na+. The latter uniformly synthesized a membrane-bound yellow pigment not found in the low salttolerant Thioalkalivibrio strains. This pigment is a 23-carbon polyene compound with a novel structure (Banciu et al. 2004b; Takaichi et al. 2004); it is clearly essential for the functioning of these bacteria at extremely high-salt and high-pH conditions. The degree of unsaturation hints on a possible antioxidant function. Under low-salt conditions, both Thioalkalimicrobium and Thioalkalivibrio 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 close to starvation ( Table 14.5) and production of different intermediates during thiosulfate oxidation suggest different pathways of sulfur metabolism in Thioalkalimicrobium and Thioalkalivibrio. The former never produced elemental sulfur during oxidation of thiosulfate, unless severe oxygen limitation was applied, while most of the Thioalkalivibrio strains formed extracellular or intracellular sulfur from thiosulfate during growth in liquid and solid culture media. Moreover, Thioalkalimicrobium was virtually unable to oxidize external elemental sulfur in contrast to Thioalkalivibrio, which converted it to sulfate. Another important difference between these two groups is sulfite metabolism. Sulfite is considered a key intermediate in sulfur oxidation pathways of many SOB species (Kappler and Dahl 2001). The Thioalkalimicrobium strains released up to 3 mM sulfite into the medium during batch growth with thiosulfate (Sorokin et al. 2000), but it was never detected in the cultures of Thioalkalivibrio. Furthermore, neither of the Thioalkalimicrobium strains studied was able to oxidize external sulfite even at micromolar concentrations, while some of the Thioalkalivibrio isolates did. Sulfite oxidation correlated with the presence of sulfite dehydrogenase activity (AMP-independent type), which was uniformly detected in aerobic strains of Thioalkalivibrio, but not in Thioalkalimicrobium (Sorokin et al. 2001a). Among the other activities, FCC type of sulfide dehydrogenase (cytochrome c-dependent), SQR (sulfide-quinone reductase), and thiosulfate reductase activities have been detected in both groups. Cytochrome c-dependent sulfide dehydrogenase was purified from Thioalkalimicrobium aerophilum AL 3 (Sorokin et al. 1998). In contrast to the classical FCC found in chemotrophic SOB (Visser et al. 1997), the enzyme from the alkaliphiles only contained cytochrome c as a cofactor. Overall, the phenotypic data suggested that in Thioalkalimicrobium, the oxidation pathway of reduced sulfur is probably similar to the one proposed for facultatively autotrophic Paracoccus species, that is, complete oxidation of the sulfane atom by a multienzyme complex (Sox) without releasing free intermediates (Friedrich et al. 2001). This is in line with the evidence for such mechanism in its close phylogenetic relative Thiomicrospira crunogena, obtained from the genome sequence (Scott et al. 2006). In contrast, Thioalkalivibrio seems to employ different mechanism(s) in the formation and consumption of free sulfur intermediates, such as elemental sulfur and sulfite, which are more common for neutrophilic SOB (Kelly et al. 1997). Currently, the complete genomes are available from a low salt-tolerant microaerophilic

Tv. sulfidophilus (Muyzer et al. 2011) and the extreme Na/K carbonate–tolerant Thioalkalivibrio K90mix (Muyzer et al. 2012). The former apparently has a duplicate sulfur-oxidizing system, including the FCC-type sulfide dehydrogenase, an incomplete Sox system and a reversed sulfidogenic system. Likewise, the latter has FCC and incomplete Sox system but lacks the rDSR. At present, it is not clear how the zero-valent sulfur atom is oxidized to the stage of sulfite in this organism. Surprisingly, despite positive results of activity measurements, both genomes lack the sqr gene. Instead, both contain the gene-encoding FCC type of sulfide dehydrogenase. This can only be explained by a possibility that the FCC (which is related to SQR) can use both cytochrome c and UQ as electron acceptors, at least in vitro. Such a possibility needs to be tested with purified FCC enzymes.

Specific Physiological Subgroups of the Soda Lake SOB Denitrification Anoxic enrichments from soda lake sediments with thiosulfate or sulfide as electron donor and nitrate as electron acceptor, even at low-salt conditions, resulted in the domination of partial denitrifiers, mainly reducing nitrate only to nitrite with copious sulfur formation. Only one enrichment culture from the hypersaline Lake Fazda in Wadi Natrun (Egypt) resulted in complete denitrification of nitrate, although with intermediate nitrite production (Sorokin et al. 2003a). Purification of this culture resulted in the selection of a stable co-culture of two facultatively anaerobic alkaliphilic SOB strains. A numerically minor population of large nonmotile coccoid cells with intracellular sulfur globules isolated in pure culture as strain ALEN2 reduced nitrate only to nitrite. Despite its obvious phenotypic difference from the known alkaliphilic SOB species (> Fig. 14.7), the 16S rDNA sequence-based phylogenetic analysis placed this unusual bacterium into the genus Thioalkalivibrio, and the strain ALEN 2 was described as a new species Tv. nitratireducens (Sorokin et al. 2003b). One of the most peculiar features of this SOB is the presence at high concentrations and with extremely high in vitro activity of a dissimilatory NiR (TvNiR), representing a novel type within the ccNiR family (Antipov et al. 2003; Tikhonova et al. 2006). Since the organism cannot reduce nitrite during anaerobic growth, the true function of the enzyme remains unclear. The genome sequence analysis of ALEN2, which is currently underway, may help to understand its function. A similar type of nitrate to nitrite-reducing SOB, strain AKLD2, has also been enriched and isolated from a Kulunda Steppe hypersaline soda lake. It had 3 % difference in 16S rRNA gene sequence with ALEN2 (see > Fig. 14.2) and remarkably unusual cells with sharp angles forming skin-like hard colonies (> Fig. 14.8). Unfortunately, the strain was lost and not fully characterized. The second (numerically dominant) organism in the denitrifying consortium, strain ALED, was a thin motile rod, which reduced the nitrite produced by its partner strain ALEN2 to N2, and grew anaerobically with

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

a

14

b

10 µm

c

d

S S

. Fig. 14.7 Cell morphology of coccoid Thioalkalivibrio. (a, c) Tv.paradoxus, (b, d) Tv.nitrati-reducens; (a, b) phase contrast, (c, d) thin sections; S intracellular sulfur globules; bars: a = 20 mm, c, d = 0.5 mm

. Fig. 14.8 Cell morphology of nitrate-reducing Thioalkalivibrio AKLD2 with unusual cell morphology (thin sections); bars: 1 mm

541

542

14

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

thiosulfate and nitrite or nitrous oxide (N2O), but not with nitrate. A similar organism, Thioalkalivibrio denitrificans ALJD, was isolated previously from a Kenyan soda lake using N2O as electron acceptor (Sorokin et al. 2001b). One of the essential properties of these SOB is the preference for N2O as an electron acceptor. In a pH-controlled continuous culture, Tv. denitrificans was able to grow anaerobically within a pH range 7.5–10.5 and an optimum at pH 9.0. Growth with N2O as electron acceptor was more stable and faster than with O2 at pH >10. Moreover, in the chemostat, it grew well with N2O and polysulfide, a form of thiodenitrification unique to alkaliphiles, since polysulfide is only chemically stable at alkaline pH and anoxic conditions. The overall reaction, performed by the thiodenitrifying consortium of two different Thioalkalivibrio species is the following: 8NO3 þ 2S2 O3 2 þ 2H2 O >>> 8NO2 þ 4SO4 2 þ 4Hþ Tv: nitratireducens 8NO2 þ 3S2 O3 2 þ 2Hþ >>> 4N2 þ 6SO4 2 þ H2 OTv: denitrificans 8NO3 þ 5S2 O3 2 þ H2 O >>> 4N2 þ 10SO4 2 þ 2Hþ Consortium

Attempts to enrich thiodenitrifying SOB from soda lakes at hypersaline conditions (2–4 M Na+, pH 10) with thiosulfate and NOx failed. Use of sulfide instead of thiosulfate, however, resulted in the selection of a high salt-tolerant Thioalkalivibrio strain ALN1 (probably a separate species most related to Tv.

a

halophilus, see > Fig. 14.2), which grew anaerobically by nitrate reduction to nitrite and oxidation of sulfide and polysulfide to elemental sulfur in saturated soda brines. At microoxic conditions, ALN1 was able to use both sulfide and thiosulfate as e-donor.

Thiocyanate Oxidation Thiocyanate (NC–S–) is a one-carbon reduced sulfur compound, which is not an easy substrate to metabolize for bacteria. Only a few neutrophilic, autotrophic SOB species were known to use it as a source of energy after first breaking it down to sulfide, ammonia, and CO2 (Kelly and Baker 1990). Aerobic enrichment cultures in a medium containing thiocyanate at pH 10 and inoculated with samples from various soda lake sediments resulted in the isolation of nine strains of haloalkaliphilic obligately autotrophic SOB growing on thiocyanate as energy and nitrogen source (Sorokin et al. 2001c). The isolates formed two distinct groups. Four motile, vibrio-shaped strains were related to the type species of the genus Thioalkalivibrio and eventually described as a new species Tv. thiocyanoxidans (Sorokin et al. 2002b, > Fig. 14.2). These strains had unusually extended periplasmic compartments when grown with thiocyanate, but not thiosulfate as energy source (> Fig. 14.9). The other five isolates

c

P

P

b

P

. Fig. 14.9 Ultrastructure of the cells of Thioalkalivibrio thiocyanoxidans ARh2 grown with thiosulfate (a) or thiocyanate (b, c) as electron donor at pH 10. P periplasm, bar 0.5 mm

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

had nonmotile, barrel-shaped cells, accumulating large amounts of intracellular sulfur, and were able to oxidize carbon disulfide (see > Fig. 14.7). Despite their obvious phenotypic difference, these strains also belonged to the genus Thioalkalivibrio, forming a separate cluster with Tv. nitratireducens (> Fig. 14.2), and were described as a new species Tv. paradoxus (Sorokin et al. 2002b). Successful enrichments with thiocyanate as substrate at pH 10 were also obtained from soda lake sediments under denitrifying conditions with nitrate—a mode of chemolithoautotrophic metabolism suggested previously by De Kruyff et al. (1957) for the neutrophilic Thiobacillus denitrificans, but never confirmed. Two closely related haloalkaliphilic SOB strains, ARhD1 and ARhD2, isolated from these enrichments were identified as members of the genus Thioalkalivibrio with closest relationship to Tv. denitrificans (> Fig. 14.2). They are described as a new species Tv. thiocyanodenitrificans (Sorokin et al. 2004b). The aerobic, thiocyanate-utilizing isolates accumulated cyanate ((N=C=O–) as an intermediate of primary thiocyanate degradation. This was the first direct confirmation of the involvement of the ‘‘cyanate pathway’’ in the primary thiocyanate degradation in pure bacterial cultures: CNS– >>> CNO– + H2S. However, in contrast to the mechanism of primary anaerobic hydrolysis of thiocyanate, resulting in the formation of cyanate and sulfide, as suggested previously (Youatt 1954; Kelly and Baker 1990), the alkaliphiles appear to employ a different mechanism of direct oxidation of the sulfane atom of thiocyanate, producing cyanate and elemental sulfur. The enzyme responsible for this action was produced in large amounts by thiocyanate-grown alkaliphilic strains and has recently been purified from the Tv. thiocyanoxidans ARh 4. A soluble monomer with an apparent molecular mass of 58 kDa, the enzyme oxidizes the sulfane atom of thiocyanate in the presence of ferricytochrome c as an electron acceptor. The protein sequence has only two to three relatives among the proteins with unknown function deposited in the database (from Methylocella, Hydrogenobacter, Thiobacillus denitrificans, and Starkeya novella with sequence homology below 50 %) (unpublished result). This indicates that the ‘‘thiocyanate dehydrogenase’’ is a novel enzyme involved in the primary thiocyanate degradation.

14

type between soda and salt lakes, where NaCl dominates, but a small fraction of sodium carbonate provides the elevated pH. A similar type is dominated in Wadi Natrun in Egypt. Strain HL17 was identified as a member of the genus Thioalkalivibrio and described as a new species Tv. halophilus. It is indeed clustering with the Wadi Natrun Thioalkalivibrio isolates demanding high Cl concentrations (> Fig. 14.2). HL17 grew within a pH range 6.5–9.8, with a broad optimum range of 8.0–9.0, thus qualifying as a facultatively alkaliphilic halophile (Banciu et al. 2004a). Another facultatively alkaliphilic halophile was isolated from hypersaline lakes with intermediate chemistry, containing both NaCl and soda approximately at the same proportions (Kulunda Steppe in southwestern Siberia, Russia, and Owens Lake in California, USA). The three strains represented a novel lineage within the group of Halothiobacillus (> Fig. 14.2) and were described as a novel genus and species Thioalkalibacter halophilus (Banciu et al. 2008). It grew up to 3.5 M Na+ within the pH range from 7.2 to 10.2 with an optimum at pH 8.5.

Culture-Independent Detection of Haloalkaliphilic SOB in Soda Lake Sediments All type strains of the haloalkaliphilic SOB isolated from soda lakes were analyzed for the presence of the cbb genes encoding the large subunit of the key enzyme (RuBisCO) of the CalvinBenson cycle of CO2 assimilation. All of them contained type Ia RuBisCO (cbbL) (Tourova et al. 2006, 2007). Using the cbbL primers updated on the basis of the sequences obtained from pure cultures of haloalkaliphilic SOB, the uncultured diversity of autotrophic bacteria in hypersaline soda lakes was investigated and compared with the results of analogous study in Mono Lake (Giri et al. 2004) and with the results of culture-dependent approach. The results demonstrated the presence of cbbL sequences related to the genus Thioalkalivibrio, but not to Thioalkalimicrobium (Kovaleva et al. 2011), while the latter was certainly present in Mono Lake. The cbbM (coding for the RuBisCO form II) sequences detected in soda lakes were much less abundant than the cbbL and only marginally identifiable as belonging to the gammaproteobacterial SOB.

Facultatively Alkaliphilic SOB from Hypersaline Alkaline Lakes

Haloalkaliphilic SOB in ‘‘Artificial Soda Lakes’’

Numerous SOB cultures enriched from soda lakes using highly buffered, sodium carbonate–based mineral medium invariably yielded obligately alkaliphilic soda-loving isolates (natronophiles) independent from Cl . On the other hand, no development was observed from the neutral salt lakes on such media. When a NaCl-based medium was used with 0.1 M NaHCO3 as buffer and carbon source at pH 9, a halophilic SOB strain was obtained from the hypersaline alkaline Stamp Lake (> Table 14.1). It was able to grow both in neutral NaCl brines and at pH values above 9 in the presence of high sodium carbonate concentrations. Stamp Lake is an intermediate lake

Thiopaq (Paques, The Netherlands) is a proven biotechnology for H2S removal from gases, based on H2S stripping from the gas phase into an alkaline solution at pH 9. The resulting ‘‘artificial soda lake’’ loaded with sulfide is a perfect medium for haloalkaliphilic SOB (Janssen et al. 2009). Indeed, recent cultureindependent and cultivation studies showed that the dominant microbial population in several of these bioreactors was represented by a microaerophilic sulfide-specialized haloalkaliphilic SOB species belonging to the genus Thioalkalivibrio and recently described as a novel species Tv. sulfidophilus (Sorokin et al. 2012). Several other alkaliphilic and alkalitolerant SOB

543

544

14

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

species belonging to the genera Thioalkalivibrio, Thioalkalimicrobium, and the Thiomicrospira pelophila group could be isolated from the reactors as well, but they usually represented minor populations. The source of these SOB species in the bioreactors (the first one was inoculated by sediments from the North Sea) is not completely clear. We suspect that there might be a dissemination of the haloalkaliphiles from deep subterranean alkaline strata (Zhang et al. 2005) throughout aquifers to the ocean. Another source of the dissemination are soda lakes themselves from which salt dust and sediment particles can be carried for long distances either by wind or by migrating water birds.

Neutrophilic Halophilic SOB from Hypersaline Habitats Chloride-Sulfate Hypersaline Habitats with Neutral pH Hypersaline aquatic habitats are divided in marine-dependent (thalassic), which include solar sea salterns, hypersaline lagoons, and deep-sea brines, and inland (athalassic) lakes formed either by evaporative concentration of incoming diluted solutions (primary evaporates) or by dissolution of ancient salt depositions (secondary evaporates). Most of the microbiology so far has been conducted on sea solar salterns. Less is known about microbial communities in hypersaline lakes, located mostly in remote areas with evaporative climate. The principal difference between these two is the much higher magnesium content in the thalassic brines and, usually, the higher sulfate content in inland lakes. Extremely halophilic, heterotrophic haloarchaea growing optimally at 3–5 M NaCl were traditionally regarded as dominating prokaryotes in these habitats (Oren 2002). Until recently, among the few extremely halophilic bacteria equal to haloarchaea in its salt response, no culturable

chemolithoautotrophs were found. The diversity of halophilic SOB able to develop optimally in NaCl brines remained largely unexplored. Apart from a single moderately halophilic species Halothiobacillus halophilus, discovered in an Australian hypersaline lake and capable of lithoautotrophic growth up to 4 M NaCl (Wood and Kelly 1991; Kelly et al. 1998; Kelly and Wood 2000), nothing was known about the true diversity of halophilic SOB. This warranted a necessity to look more closely at this group in hypersaline chloride-sulfate habitats with neutral pH. The exploration of these habitats demonstrated unexpectedly high culturable diversity of halophilic SOB and resulted in description of seven novel SOB taxons (Sorokin et al. 2006a; Sorokin 2008). Hypersaline aquatic habitats in six different regions have been examined during 2002–2008, including hypersaline inland lakes at four sites, a sea solar saltern, and a deep-sea salt brine (> Table 14.6). The main area of study was the Kulunda Steppe, southwestern Siberia, located along the northeastern Kazakhstan border. It contains numerous salt lakes with a total salt content from 10 % to 38 % (w/v), a pH range from 7.5 to 8.5, and with Na+, Mg2+, Cl , and SO42 as the dominant ions in the brines. A typical lake in this area is shown in > Fig. 14.10. Other lake provinces were studied only briefly. In addition, a sample from a final evaporation pond in Secˇovlje Adriatic Sea saltern (Gunde-Cimerman et al. 2000) and a sample of deep-sea brine from the eastern Mediterranean Urania Basin (van der Wielen et al. 2005) were included into the analysis as examples of thalassic habitats.

Enrichment of Halophilic SOB Enrichment and subsequent cultivation of halophilic SOB was performed on a mineral medium containing either 2 M (for moderate halophiles) or 4 M (for extreme halophiles)

. Table 14.6 Investigated hypersaline habitats with neutral pH Mineral composition Salinity (g l 1)

Na+ (M)

Na+/ Mg2+

Cl (M)

Cl / SO42

7.5– 8.5

100–380

0.9– 5.0

14

0.65– 4.4

4.5

Lakes Dzun-Davst and Dolon- 7.5– Davst 8.1

220–320

No data

Region

Type

Name

pH

Kulunda Steppe (Southwestern Siberia)

Inland lakes

20 small lakes

Northeastern Mongolia South Russia

Lake Baskunchak

6.2

360

Crimean peninsula (Ukraine)

Lakes Marfinskoe and Kayashskoe

8.0

115–145

Slovenia (Adriatic coast)

Sea solar saltern

Secˇovlje

6.5

320

4.1

5.2

5.2

18.6

East Mediterranean

Deep-sea brine

Urania Basin

6.8

220

3.10

11.1

3.30

34.7

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

. Fig. 14.10 A typical hypesaline lake in Kulunda Steppe (Atlai, Russia)

NaCl. The mineral base also contained 1–2 g/L K2HPO4 and 5 mM (NH4)2SO4, pH 7.0. After sterilization, the medium was supplied with trace metals and 2 mM Mg sulfate. NaHCO3 (added from a filter-sterilized stock with pH 8.0) served as a carbon source and additional alkaline buffer (pH 7–8). To prevent loss of CO2 and evaporation, aerobic cultivation was performed in closed bottles with 10 % liquid volume at static conditions. The pH monitoring and maintaining between 7 and 7.5 was crucial during the cultivation, and it was achieved either by adding extra NaHCO3 or by injecting sterile CO2 into the gas phase. In most cases, thiosulfate (10–20 mM) was used as the energy source and, in some cases, also sulfide, tetrathionate (5 mM), or thiocyanate (10 mM). The gradient cultivation technique with sulfide (Nelson et al. 1986) in 25 ml cylinders was employed. Solid medium containing 2–3 M NaCl was prepared by mixing complete liquid medium containing 4 M NaCl and 30–40 mM thiosulfate with 4–6 % (w/v) washed agar at different ratio at 50oC. The plates were incubated in closed jars at 0–20 % O2/5 % (v/v) CO2 in the gas phase. The enrichment strategy and the general isolation results from different enrichments from hypersaline habitats are shown on > Fig. 14.11 and summarized in > Tables 14.7 and > 14.8.

Moderately Halophilic Aerobic SOB Aerobic enrichments in dilutions with thiosulfate at 2 M NaCl inoculated by sediments from hypersaline lakes developed quite rapidly and were positive up to 10 5–10 7 dilution. At full aeration, short motile rods forming large sulfur-containing colonies were the dominant morphotype, while in static cultures with a high liquid/gas ratio and with sulfide as substrate in gradient enrichments, a highly motile small vibrio became the dominant morphotype. It also could produce tiny sulfur colonies on thiosulfate plates after prolonged incubation. After isolation in pure culture, the vibrio strains could easily grow in fully aerated culture. Overall, four rod-shaped (> Fig. 14.12a)

14

and three vibrio-shaped strains (> Fig. 14.12b) were obtained in pure culture from Siberian and Mongolian lakes (> Table 14.7). The rod-shaped isolates were closely related to Halothiobacillus halophilus, while the vibrio strains clustered with members of the Thiomicrospira crunogena group and were described as a novel species Tm. halophila (Sorokin et al. 2006a, b) (> Fig. 14.13). In the enrichment from the deep-sea Urania brines, only rod-shaped phenotype was present, and an isolate strain HL-U1 was closely related to Halothiobacillus hydrothermalis (> Fig. 14.13). Two more Halothiobacillus strains, HL20 and HL27, were isolated from the aerobic enrichments at 4 M NaCl, when highly specialized extremely halophilic SOB (see below) were either absent or present at low numbers. Strain HL20 dominated an enrichment culture from a Mongolian lake with tetrathionate as substrate, while strain HL27 was one of the dominant organisms in the enrichment culture from a Crimean lake with thiosulfate. All these isolates were moderately halophilic growing up to 3–4 M NaCl, but with a much lower optimum at 0.5–1 M NaCl, similar to Htb. halophilus (> Fig. 14.14).

Moderately Halophilic Thiodenitrifying SOB Anaerobic enrichments at 2 M NaCl with thiosulfate as e-donor and nitrate as acceptor were positive in several enrichments from lake sediments and from a saltern. Although nitrite, N2O, and elemental sulfur were copiously released as major nitrogen and sulfur intermediates, in contrast to the soda lake enrichments, the denitrifying cultures from hypersaline neutral habitats produced N2 and sulfate as final products, and single organisms were responsible for the whole process. The facultative anaerobic isolates thus obtained were represented by long, non-motile, rod-shaped cells (> Fig. 14.12f). Genetically, the isolates formed a single species and formed a new lineage within the Gammaproteobacteria with the closest relatives among a cluster of marine unclassified thiodenitrifyers (Nercessian et al. 2005; Sievert and Muyzer, unpublished) (> Fig. 14.13, > Table 14.7). The group was described as a new genus and species Thiohalomonas denitrificans (Sorokin et al. 2007a). Despite the fact that washed cells, grown with nitrate, could reduce nitrite and N2O to N2 in presence of thiosulfate, the anaerobic growth occurred only with nitrate. Growth was also possible under microoxic conditions at O2 concentration below 5 % (v/v) in the gas phase. The Thiohalomonas strains are moderate halophiles with a relatively narrow salt range for growth (both aerobic and anaerobic) between 1 and 3 M NaCl (> Fig. 14.14).

Extremely Halophilic Aerobic SOB Aerobic enrichment cultures at 4 M NaCl developed much slower than at 2 M, and there was a major problem with their pH maintenance: during the initial stage, the brines gradually became alkaline, and periodic injection of CO2 was necessary to

545

546

14

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

Sediments from hypersaline lakes, sea saltern, deep-sea brines

Enrichments at 2 M NaCl

Fully aerobic, colonies

S2O32−

Halothiobacillus

NCS−

Thiohalobacter

Enrichments at 4 M NaCl

Anaerobic denitrifying

Microaerophilic

Thiomicrospira halophila

S2O32−/ NO3−

Thiohalomonas

Aerobic, microaerophilic

NCS−/ NO3−

Thiohalospira

Anaerobic denitrifying

Thiohalorhabdus

Thiohalophilus

. Fig. 14.11 Generalized scheme of enrichment strategy and isolation of halophilic SOB from hypersaline habitats with neutral pH

keep the pH below 8, while on the late stages, addition of NaHCO3 became necessary. After solving the pH problem, positive enrichments were obtained for most of the samples studied except the deep-sea brines. This indicated universal presence in the hypersaline habitats of SOB populations able to develop at saturating salt concentrations. Moreover, they were as abundant in the lake sediments as the moderate halophiles (103–107 per cm3). The dominant phenotype observed at 4 M NaCl in most cases was a thin motile spirillum (> Fig. 14.12c). Overall, 20 SOB strains of this type were obtained from salt lakes and a saltern with either thiosulfate, sulfide, or tetrathionate as substrates (> Table 14.7). Based on 16 S rRNA gene sequence analysis, it represents a new lineage in the Gammaproteobacteria clustering with the members of the family Ectothiorhodospiraceae (> Fig. 14.13). The group was described as a new genus and species Thiohalospira halophila. All strains are extreme halophiles growing between 2 and 5 M NaCl—an ecotype not known before among any described chemolithoautotrophs (> Fig. 14.14). Another specific property of this group was production of large amounts of tetrathionate as an intermediate of thiosulfate oxidation (up to 80 % conversion), which was finally oxidized to sulfate. A second species to this genus, Thsp. alkaliphila, was found in hypersaline lakes with alkaline pH (Wadi Natrun). Although it was enriched on the standard medium for halophilic SOB at 2 M NaCl, strain ALgr6sp was able to grow up to pH 10 and is qualified as facultatively alkaliphilic halophile (Sorokin et al. 2008a).

Extremely Halophilic Denitrifying SOB Positive anaerobic enrichments with thiosulfate and nitrate at 4 M NaCl were obtained from four lake samples and from a saltern. They developed very slowly, but, due to the very selective condition, there were very few heterotrophic satellites, and all five positive enrichments yielded a pure culture of the dominant SOB morphotype with long flexible nonmotile, rod-shaped cells (> Fig. 14.12e). A similar phenotype was also found in two aerobic enrichments at 4 M NaCl. Strain HL19 was isolated from a mixotrophic enrichment with acetate/thiosulfate, in which it developed on tetrathionate produced from thiosulfate by an acetate-utilizing haloarchaneon (Sorokin et al. 2005). Despite being enriched at aerobic conditions, similar to the HLD strains, it was able to grow anaerobically with nitrate. Another similar strain, HL 28, was isolated from an aerobic enrichment culture at 4 M NaCl from one of the Crimean lakes, where it was developing in a mixture with a Thiohalospira (see above). All isolates were related at the species level and formed a new deep lineage within the Gammaproteobacteria most probably of a family level, only distantly related to the genus Acidithiobacillus (> Fig. 14.13). The group was described as a new genus and species Thiohalorhabdus denitrificans (Sorokin et al. 2008b) (> Table 14.7). Both aerobic and anaerobic cultures, similar to Thiohalospira, could not grow below 2 M NaCl and had an optimum at 3 M, thus belonging to the extreme halophiles (> Fig. 14.14).

1

10

MhA SCN

MhD

SL, ST

20

7

EhA

EhD

SL, ST

SL

MhD, SCN 1

SL, ST

SL

SL

Thiohalorhabdus

Thiohalospira

Thiohalophilus

Thiohalomonas

Thiohalobacter

Thiomicrospira crunogena –

–

2.0–4.5 (3.0)

2.0–5.0 (3.0)

1.0–4.0 (2.5) +

–

+

1.0–3.0 (1.5–2.0) +

0.2–3.0 (0.5)

0.5–3.5 (1.5)

0.5–4.0 (1.0–1.5) –

S4O62

Sulfur

Denitrification S-intermediate

–

–

+

–

+

–

–

Ia, II

Ia, II

Ia

Ia, II

Ia

Ia

Ia, II

65.0–65.8

65.8–67.0

58.2

58.0–60.0

63.5

56.1–57.1

64.0–67.7

b

1

3

1

2

1

2

4.4

3.5

5.6 0.052

4.2

0.03–0.04 2.0–2.5

0.10

0.03–0.04 4.0–4.9

0.045

0.25

0.20–0.35 4.0–4.5

Y

Growth kineticsd

GeneCNS oxidation RuBisCO type G + C mol% speciesc m

Genetic properties

MhA moderately halophilic aerobes, MhD moderately halophilic denitrifiers, SCN thiocyanate-utilizing, EhA extremely halophilic aerobes, EhD extremely halophilic denitrifiers SL inland lakes, MB deep-sea brines ST, sea salterns c On the basis of DNA-DNA hybridization d With thiosulfate : m specific growth rate at optimal salinity (h 1), Y specific growth yield (mg protein mmol 1). The new genera are in bold

a

3

MhA

Halothiobacillus halophilus

7

MhA

SL, MB

Number of isolates Habitatb Affiliation

Typea

Salt range (optimum) M NaCl

. Table 14.7 Culturable halophilic SOB types in hypersaline habitats: summary

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

14 547

548

14

Halophilic and Haloalkaliphilic Sulfur-Oxidizing Bacteria

. Table 14.8 Respiratory activity of washed cells of haloalkaliphilic SOB V, mmol O2 (mg protein min) Organism

No of tested strains

NaCl, M

HS

S2O32

Halothiobacillus sp.

4

2.0

1.8–3.2

1.0–1.5

1

S4O62

S8

NCS

0–0.1

0.2–0.4

0

Thiomicrospira halophila

3

1.5

0.9–1.2

0.5–0.6

0

0.08–0.12

0

Thiohalomonas denitrificans

5

2.0

0.1–0.2

0.15–0.25

0.08–0.10

0.08–0.1

0

Thiohalobacter thiocyanaticus

1

1.0

0.48

0.35

0.2

0.24

0.08

Thiohalophilus thiocyanoxidans

1

2.0

0.36

0.28

0.06

0.04

0.11

Thiohalospira halophila

6

3.0

0.25–0.80

0.38–0.62

0.13–0.34

0.07–0.30

0

Thiohalorhabdus denitrificans

3

3.0

0.14–0.48

0.15–0.50

0.10–0.28

0.06–0.10

0

Abbreviations: V respiration rate, mmol of O2 (mg of protein min) 1; N number of tested strains

In contrast to the moderately halophilic denitrifying Thiohalomonas strains (see above), the extremely halophilic thiodenitrifyers grew well under microoxic conditions (2–5 % (v/v) oxygen) and some of them even in fully aerated cultures. Tetrathionate was a major intermediate of aerobic thiosulfate oxidation to sulfate in this group, similar to the aerobic extreme halophiles from the genus Thiohalospira. This fact tempts us to speculate that tetrathionate formation as a major intermediate is somehow connected with the growth of SOB at extreme salinity. In both Thiohalospira and Thiohalorhabdus isolates, a very active periplasmic tetrathionate synthase has been detected optimally functioning at 3–4 M salt and with a clear demand for both sodium and chloride ions. Under anaerobic growth conditions, with either thiosulfate or tetrathionate as substrates, nitrate was only reduced to nitrite, and sulfur accumulated as an intermediate. However, washed cells, grown with nitrate, were capable of slowly reducing nitrite and, more actively, N2O in the presence of thiosulfate as electron donor. Therefore, restriction of anaerobic growth to nitrite formation is most probably the result of inhibitory effect of saturated salt concentration on periplasmic nitrite and N2O reductases.

Halophilic Thiocyanate-Oxidizing SOB So far, there was only a single report on thiocyanate oxidation in industrial saline wastewater (salinity up to 30 g/L), where the responsible SOB was identified as a member of the genus Halothiobacillus (Stott et al. 2001). Positive enrichments with thiocyanate as energy and nitrogen source from hypersaline habitats were obtained only at moderate salinity, that is, 1–2 M NaCl. At aerobic conditions, several identical strains of moderately halophilic chemolithoautotrophic SOB, utilizing thiocyanate as e-donor, were isolated. One of these isolates, strain HRh1, (> Fig. 14.13) had been characterized in detail and described as a novel genus and species Thiohalobacter thiocyanaticus within the Gammaproteobacteria (Sorokin et al. 2010) (> Fig. 14.12d). It grew with thiocyanate as the only substrate at salinity up to 2 M NaCl and with thiosulfate—up

to 3 M NaCl with an optimum at 0.5 M (> Fig. 14.14), thus being moderately halophilic. The isolate accumulated cyanate as an intermediate of the primary thiocyanate degradation, similar to the thiocyanate-utilizing Thioalkalivibrio strains from soda lakes, but the overall activity of thiocyanate oxidation in the halophilic strain HRh1 was much lower than in the soda lake isolates. The thiocyanate-growing cells overexpressed a single soluble protein with an apparent molecular mass of 60 kDa, which was absent in the cells grown with thiosulfate. Under anaerobic conditions with thiocyanate as electron donor and nitrate as electron acceptor at 2 M NaCl, a single stable binary culture was selected from a mixture of hypersaline lake sediments, which eventually resulted in the isolation of strain HRhD2 capable of aerobic growth with thiocyanate as the only substrate (> Fig. 14.12g). Phylogenetic analysis of strain HRhD2 placed it to a novel lineage within the Gammaproteobacteria (> Fig. 14.13) and was described as a new genus and species Thiohalophilus thiocyanatoxidans (Sorokin et al. 2007b). Aerobic growth with thiocyanate was possible from 1 to 4 M NaCl with an optimum at 1.5 M (> Fig. 14.14), and, so far, this organism is representing the most halophilic SOB growing with thiocyanate. The final products of thiocyanate metabolism were sulfate and ammonium. In contrast to Thiohalobacter and Thioalkalivibrio, Thiohalophilus degraded thiocyanate via the ‘‘COS’’ pathway using the enzyme thiocyanate hydrolase (Katayama et al. 1998), which was purified and characterized (Bezsudnova et al. 2007). Despite the absence of cyanate among the intermediates of thiocyanate degradation in Thiohalophilus, a high activity of the enzyme cyanase had been detected in the cells, grown with thiocyanate, but not with thiosulfate. This fact once more indicated that the pathway of thiocyanate degradation in SOB cannot be elucidated on the basis of the presence of this secondary enzyme; that is, the presence of cyanase does not necessarily mean that thiocyanate degradation is going through cyanate, as had been suggested previously (Youatt 1954). The bacterium was also able to grow anaerobically with thiosulfate using nitrite (but not nitrate) as the electron acceptor at low concentrations ( Table 15.1a, which shows many of the genera identified as obligately chemolithoautotrophic colorless sulfur bacteria, because their primary metabolism involves the oxidation of inorganic energy sources and the fixation of CO2 (or CO). The oxidation of reduced sulfur compounds is a major feature of their metabolism, but many of them can also use other substrates such as H2 and transition metals. > Table 15.1b shows genera that contain more versatile species which can grow heterotrophically and can also either grow chemolithoautotrophically or chemolithoheterotrophically on reduced sulfur compounds. There are other species such as Catenococcus thiocyclus (Sorokin 1992; Sorokin et al. 1996), which seem to gain a small amount of supplemental energy from one of the oxidation steps, or may use one of the reactions to detoxify a metabolite (e.g., sulfide). Over the last few years, there have also been reports that mitochondria from cells ranging from the gills of mussels to human guts are also able to generate energy from sulfide oxidation (Searcy 2006; Goubem et al 2007; Wendeberg et al. 2012). The ability to gain energy from the oxidation of reduced sulfur compounds is clearly widespread and has little taxonomic significance. That said, in ecological terms, the ability is important, and it is convenient to consider these bacteria as a group. The colorless sulfur bacteria play an essential role in the oxidative side of the sulfur cycle (> Fig. 15.1). Like all of the element cycles, the sulfur cycle has oxidative and reductive sides, which, in most ecosystems, are in balance. However, this balance does not always exist, and accumulations of intermediates such as sulfur, metallic 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. 15.1). The adjective ‘‘colorless’’ is used because of the lack of photopigments in these bacteria, although it should be noted that colonies and dense cultures could 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 elsewhere. Taxonomic details about the various genera shown in > Table 15.1a and b can be found in their respective chapters elsewhere in this handbook. As mentioned above, the ability to grown on reduced sulfur compounds was once regarded as taxonomically significant. In 2000, Kelly and Wood (2000) rationalized a situation in which obligate and facultative autotrophs, acidophiles and neutrophiles, and thermophiles and psychrophiles had all previously been included in the same genera, depending on the morphology of the cells involved (e.g., Thiobacillus, Thiomicrospira, Thiosphaera). They took into account the physiology and phylogeny of the various species and devised new

names for the various genera that provided a bit more information about the species therein. For readers more accustomed to the historical names, > Table 15.2 summarizes the changes to the names of the older colorless sulfur bacteria. To avoid confusion, the modern names will be used throughout this chapter. 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, ferrous iron, or even arsenic compounds. As well as differences in substrate range, there is also some variation in electron acceptor usage. Although most colorless sulfur-oxidizing bacteria require oxygen, some are able to grow anaerobically using nitrogen oxides (e.g., nitrate) as their terminal electron acceptor during denitrification. Others have been shown to use other oxides such as arsenate. 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 serves as electron donors. Acidithiobacillus ferrooxidans can 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 15.3. In the following sections, we will first discuss the physiology of the colorless sulfur bacteria and then cover taxonomic aspects. This will be followed by a discussion of the habitats of the colorless sulfur bacteria, including artificial habitats, and finally some ways in which they are used. 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 techniques available for the measurement of their activities.

Physiology The great diversity of colorless sulfur bacteria should come as no surprise if it is remembered that the group encompasses Archaea as well as Bacteria, and that the latter group is also very diverse, including common pseudomonads and organisms that might be considered as ‘‘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 that can be grown in the laboratory. This is particularly true of our understanding of the biochemistry of sulfur metabolism and, to a lesser extent, of carbon metabolism.

Colorless Sulfur Bacteria

15

. Table 15.1 (a) Obligately chemolithoautotrophic genera that can obtain energy for growth from one or more reduced sulfur compounds. Many can also use other substrates such as H2 and transition metals such as Fe(II). (b) Facultative chemolithoautotrophs and others able to gain energy from oxidizing reduced sulfur compounds Publication establishing genus

Group

Optimum temperature  C pH Optima

Source

Other

Sulfuricella

b

22

7.5–8.0

Freshwater lake

Denitrifies NO3 to N2

Kojima and Fukui (2010)

Thiobacillus

b

28–43

2.0–8.0

Ubiquitous

Some denitrify, others do not

Beijerinck (1904), Kelly and Wood (2000)

Thiobacter

b

50–55

6.5–7.0

Geothermal aquifer

Acidithiobacillus

g

25–45

2.0–3.5

Acid-mine drainage, Can grow with Fe(II) as sole Kelly and Wood sulfidic leachate energy source (2000)

Halothiobacillus

g

28–40

6.5–8.0

Seawater at Naples

Halophilic, 0.4–1M NaCl

Kelly and Wood (2000)

Sulfurivirga

g

50–55

6.0

Hydrothermal microbial mat

Uses thiosulfate and tetrathionate

Takai et al. (2006)

Thioalkalibacter

g

30

88.0–10.2

Soda lakes

Optimum Na+ concentration 1.0 M

Banciu et al. (2008)

Thioalkalimicrobium

g

Mesophilic

10.0

kenyan soda lakes

Grows at 0.2–4 M NaCl, some species denitrify

Sorokin et al. (2001)

Thioalkalispira

g

30

10

Egyptian soda lake

Microaerobic, NO3 only reduced to NO2 , NaCl optimum 0.5 M

Sorokin et al. (2002)

Thioalkalivibrio

g

40

10.0–10.2

Kenyan soda lakes

Requires NaCl, 0.3–4 M depending on species. carboxysomes

Sorokin et al. (2001)

Thiofaba

g

45

6.5

Hot springs

Thiohalobacter

g

32

7.3–7.5

Hypersaline chloride-sulfate lakes

Can use SCN as substrate, Sorokin et al. (2010) requires NaCl (optimum 0.5 M)

Thiohalomonas

g

Mesophilic

7.3–8.2

Hypersaline lakes

Optimum NaCl 1.5–2.0M, denitrify, microaerobic

Thiohalophilus

g

32

7.3–7.5

Hypersaline chloride-sulfate lakes

NaCl optimum 0.5 M, can Sorokin et al. (2007) use SCN as energy source

Thiohalorhabdus

g

33–35

7.5–7.8

Hypersaline lake sediments

NaCl optimum 3 M

Thiohalospira

g

32–35

7.3–7.8

Hypersaline habitats NaCl optimum 3 M, copious S4O62 produced during S2O32 oxidation

Sorokin et al. (2008)

Thiomicrospira

g

28–30

6.5–8.0

Hydrothermal mud

Req NaCl up to 3 %

Kuenen and Veldkamp (1972)

Thioprofundum

g

50

7.0

Deep sea hydrothermal vents

Anaerobic-microaerobic, NO3 to N2 and N2O

Takai et al. (2009)

Thiovirga

g

30–34

7.5

Wastewater treatment biofilm

Opt. NaCl 18 mM; max NaCl 180 mM carboxysomes

Ito et al. (2005)

Sulfuricurvum

2

10–35

7.0

Crude oil storage cavity

Denitrifies NO3 , not NO2 , microaerobic

Kodama and Watanabe (2004)

Genus name (a)

Hirayama et al. (2005)

Mori and Suziki (2008)

Sorokin et al. (2007)

Sorokin et al. (2008)

557

558

15

Colorless Sulfur Bacteria

. Table 15.1 (continued) Source

Other

Publication establishing genus

6.14

Hydrothermal, polychaetes’ nest

Denitrifies

Inagaki et al. 2003

6.5–7.0

Hydrothermal vent sediment

Denitrifies

Inagaki et al. (2004)

Sulfide/oxygen interface in water

Chemotactic for O2 and H2S, characteristic veils, chemotactic

Hinze (1913)

7.0

Soil

Requires additives such as Kelly et al. (2000) biotin, autotrophic growth on formate

35

8.0

Near-shore hydrothermal

Optimum NaCl 35 g/l

Sorokin et al. (2005)

b

20–30

6.5–7.5

Contaminated running water, activated sludge

Sheathed filaments

Kutzing (1833)

Sulfuritalea

b

25

6.7–6.9

Japanese freshwater Autotrophic growth only lake anaerobically, denitrifies. H2 also energy source

Beggiatoa

g

25–38

Neutrophilic Sulfide/oxygen interface in sea and freshwater

Stores NO3 for denitrification, can reduce intracellular So, filamentous

Thermithiobacillus

g

43–45

6.8–7.5

Heterotrophic growth on Kelly and Wood complex media, not simple (2000) substrates

Thiomargarita

g

Moderate psychrophile

Neutrophilic Sulfidic marine Cells 100–200 mm, stores sediments off South NO3 and S2 in an America intracellular vacuole, sheathed, not in pure culture

Schulz et al. (1999)

Thioploca

g

Moderate psychrophile

Neutrophilic Oxic/anoxic interface

Filamentous in a twisted braid, not in pure culture, gliding motility, stores NO3 and S2 in an intracellular vacuole

Lauterborn (1907)

Thiothrix

g

25–30

6.5–8.5

Sulfur springs and activated sludge plants

Filamentous, sheathed, may produce rosettes

Winogradsky (1888)

Acidianus

Archaea

90

2.0

Solfataras and marine hydrothermal vent systems

Aerobic So oxidation, anaerobic So reduction with H2

Segerer et al. (1986)

Sulfurisphaera

Archaea

84

2.0

Acid hot springs

Anaerobic growth with So as e acceptor

Kurosawa et al. (1998)

Sulfolobus

Archaea

75–80

2.0–3.0

Volcanic springs

Heterotrophic growth only Brock et al. (1972) with O2, anaerobic growth with So as e- acceptor

Sulfurococcus

Archaea

40–80

Acidophilic

Hydrothermal

Also uses ferrous

Golovacheva et al. (1995)

7.5

Hot aquifers

Some species use H2

Takai et al. (2003)

Genus name

Group

Optimum temperature  C pH Optima

Sulfurimonas

2

.30

Sulfurovum

2

28–30

Thiovulum

2

Fig. 15.2a, b). Not only do the enzymes and electron carriers differ but also their location 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 to drive the Calvin cycle. In most obligate and facultative autotrophs, the Calvin cycle serves as the route for carbon dioxide fixation. 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 and energy metabolism. > Table 15.4 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 certain 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 Archaea [which use a reductive carboxylic cycle (Ko¨nig 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 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 secondary metabolic activities, the organisms being primarily dependent on a lithotrophic energy source and carbon dioxide for autotrophic growth. > Table 15.1a shows the genera that contain species which fall into this group.

Thiobacillus capsulatus

Facultative Chemolithotrophs Although the biochemistry of the oxidation of sulfur compounds received much attention during the twentieth century, the pathways involved were not well understood. This was due, in particular, to the fact that the research was formulated around

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

Colorless Sulfur Bacteria

. Table 15.3 Examples of the reactions used by the colorless sulfur bacteria to gain energy for growth H2 S þ 2O2 ! H2 SO4 2H2 S þ O2 ! 2S0 þ 2H2 O 2S0 þ 3O2 þ 2H2 O ! 2H2 SO4 Na2 S2 O3 þ 2O2 þ H2 O ! Na2 SO4 þ H2 SO4 4Na2 S2 O3 þ O2 þ 2H2 O ! 2NaS2 O3 þ 4NaOH 2Na2 S4 O6 þ 7O2 þ 6H2 O ! 2Na2 SO4 þ 6H2 SO4 2KSCN þ 4O2 þ 4H2 O ! ðNH4 Þ2 SO4 þ K2 SO4 þ 2CO2 5H2 S þ 8KNO3 ! 4K2 SO4 þ 4N2 þ 4H2 O

15

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, Thiobacterium, 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).

5S0 þ 6KNO3 þ 2H2 O ! 3K2 SO4 þ 2H2 SO4 þ 3N2

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 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 requiring (partially) separate metabolic pathways or enzymes where diauxie or biphasic growth might occur 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 15.1b lists genera that contain species able to grow on mixtures of reduced sulfur compounds and organic substrates. To some extent, the phototrophic sulfur-oxidizing 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 group of bacteria is characterized by an ability to generate energy from the oxidation of reduced sulfur compounds but which cannot fix carbon dioxide. It is not always obvious whether a species fits into this group as specialized growth conditions may be required both to establish that energy is generated and to exclude autotrophy. For example, for a long time, Thiomonas perometabolis was considered to be chemolithoheterotrophic but was then shown that under certain conditions, it can grow autotrophically (Katayama-Fujimura et al. 1984). However, other chemolithoheterotrophic species have been isolated, and a few strains have been well characterized (e.g., Tuttle et al. 1974; Gommers and Kuenen 1988). Some Beggiatoa strains may belong in this group (Larkin and Strohl 1983). As is clear from the example of Thiobacillus perometabolis, careful testing under a variety of conditions is necessary in order to discriminate chemolithoheterotrophs from the facultative autotrophs and the sulfur-oxidizing heterotrophs.

Electron Acceptors for Aerobic and Anaerobic Growth Oxygen is almost 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. 15.3. 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. The ability to denitrify is not limited to any particular physiological type (see > Table 15.1a, b). A few species can only use part of the nitrification pathway. For example, Thiobacillus thioparus can only reduce nitrate to nitrite and requires the presence of a nitrite-reducing bacterium for anaerobic growth. Strictly speaking, of course, the latter reaction is not truly denitrification, but since the reaction still serves for electron transport and survival under anaerobic conditions, these species are appropriately included here. A few obligately chemolithotrophic sulfur bacteria 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). Sulfurimonas 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. For example, the facultative chemolithotroph Paracoccus pantotrophus has been found to retain its sulfur-oxidizing 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 Sulfurimonas denitrificans (0.06 h 1). Many other facultatively autotrophic bacteria lose their sulfur-oxidizing capacity in anaerobic cultures but are still able to denitrify using organic compounds, or even

561

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Colorless Sulfur Bacteria

a −S − SO3−

SH2O

ENZYME A (Mr 16,000) Binds thiosulphate [A – SO3 – S –] in a 1:1 molar ratio ENZYME B Mr 63,000; subunits.Mr 32,000; contains Mn in a 1:1 molar ratio 2SO42− 10H+

Sulphite: cytochrome c oxidoreductase (Mr 44,000) Cytochrome Css Mr 260,000; aggregate of six polypeptides of Mr 43,000; 4–5 haem and 6–7 Fe per mole; Two mid-point redox potential centres, Em,7–115 and +240 mV

8H+ + 2O2

cytochrome oxidase (aa3)

4H2O

8e−

cytochrome Css2 8e−

Cytochrome Css2.5 Mr 56,000;subunits Mr 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

Outre membrane

Cytoplasmic membrane

Periplasm

b

2−O3-S-SO3− trithionate 2 H2 O

2 SO42−

trithionate hydrolase 4 H+ inhibition by FCCP

2S2O2− tetrathionate 3 synthose

S4O42−

S4O42− 2 e−

cytochrome oxidose

(Mechanism unkown) 4 SO32−

cytochrome c

8e−

Inhibition by HONO

4 H2O sulfite dehydrogenase 8 H+

Ubiquinone cytochrome b 4 SO42−

Outer membrane

. Fig. 15.2 (Continued)

Periplasmic space

4 SO42−

Cytoplasmic membrane

Cytoplasm

Colorless Sulfur Bacteria

hydrogen. Among these are Paracoccus versutus and Paracoccus denitrificans (Taylor and Hoare 1969; Friedrich and Mitrenga 1981). Sulfide-dependent reduction of nitrate to N2 by Beggiatoa tufts was shown using 15N-labeled nitrate (Sweerts et al. 1990). Members of the genus Sulfuritalea are unusual in that they can only grow autotrophically under anaerobic conditions, when they denitrify (Kojima and Fukui 2011). Among the sulfur-oxidizing genera within the Archaea, Sulfolobus species appear to be the most dependent on oxygen, although some have been shown to use ferric iron or molybdate as electron acceptors under microaerobic conditions (Brock and Gustafson 1976; Brierley 1982). Members of the genera Sulfurisphaera and Acidianus are among those able to grow under anaerobic conditions, by using sulfur as their electron acceptor, thus making these bacteria both sulfuroxidizing and sulfur-reducing, depending on the conditions (Segerer and Stetter 1989; Kurosawa et al. 1998). Nelson and Castenholz (1981) 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 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, Halothiobacillus 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, Thiobacillus ferrooxidans can use ferric iron as an electron acceptor.

Ecophysiology as a Function of pH, Temperature, and Nutrient Availability Colorless sulfur bacteria have been found growing at pH 1.0 and pH 11.0, at 4  C and 95  C, and at dissolved oxygen

15

concentrations ranging from air-saturation to anaerobiosis (> Table 15.1a, b). 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 genera of colorless sulfur bacteria are shown in > Table 15.1a, b. 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 thiosulfatelimited 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 P. versutus and Thiobacillus 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. Many of them have specialized in growth in the gradients where (anaerobic) sulfide-containing 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. This group includes mesophilic obligate and facultative autotrophs (e.g., Acidithiobacillus ferrooxidans and Acidithiobacillus 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, Acidithiobacillus ferrooxidans is able to grow mixotrophically on the iron and sulfur components of pyrite (Arkestein 1980) or on mixtures of ferrous iron and

. Fig. 15.2 Pathways of oxidation of reduced sulfur compounds in two different organisms. (a) The periplasmic thiosulfate-oxidizing system of Paracoccus 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 Thermithiobacillus tepidarius as proposed by Kelly (1988b). In contrast to the system shown in (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 carbonyl cyanide-ptrifluoromethoxyphenylhydrazone, HQNO 2- heptyl-4-quinolinol-1-oxide

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. Table 15.4 The different physiological types of colorless sulfur bacteriaa Carbon source Physiological type

Inorganic

Obligate chemolithotroph

+

Facultative chemolithotroph

+

Energy source Organic

Inorganic

b

Organic

+ +

+

+

Chemolithoheterotroph

+

+

+

Heterotroph

+

+

a

Commonly used synonyms for chemolithotroph include chemolithoautotroph, autotroph, chemoautotroph, and lithotroph +, used by the group; -, not used by the group

b

-

0

+ pO2 (ell) Sulfurimonas denitrificans Thiobacillus denitrificans Thiomicrospira pelophila Thiobacillus thioparus Acidithiobacillus thiooxidans

. Fig. 15.3 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 (Based on Timmer ten Hoor 1977)

tetrathionate, gaining energy from the iron- and sulfuroxidizing reactions (Hazeu et al. 1986, 1988). There were a few reports of facultatively heterotrophic growth by Acidithiobacillus ferrooxidans (e.g., Shafia and Wilkinson 1969; Lundgren et al. 1964). However, it has since been shown that most of the Acidithiobacillus ferrooxidans cultures available from culture collections were contaminated with acidophilic facultative autotrophs and heterotrophs (Harrison 1984), including Acidithiobacillus acidophilus and Acidiphilium cryptum, and it is now generally accepted that Acidithiobacillus ferrooxidans is an obligate autotroph. It has frequently been assumed that Acidithiobacillus 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 Acidithiobacillus ferrooxidans for an immunofluorescent assay of slurries made from coal from different sources. Unsterilized and sterilized coal samples were inoculated with Acidithiobacillus ferrooxidans, with a mixed culture of pyriteoxidizing bacteria from a coal-washing installation, and a mixture of the two. Despite the fact that a DNA-fluorescent stain indicated abundant microbial life in all of the slurries, the only sample in which a significant Acidithiobacillus ferrooxidans population was detected was the control, which had been sterilized and then inoculated with the pure culture of Acidithiobacillus ferrooxidans. It appears that in all other cases, other strains (which might include such species as Acidithiobacillus thiooxidans, Leptospirillum ferrooxidans, or Acidiphilium cryptum, to name but a few) were able to successfully outcompete Acidithiobacillus 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  C to 95  C. However, the majority of the frequently studied species are mesophilic. 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). The bacteria in this group can be subdivided into two groups, the moderate thermophiles (generally Bacteria), which grow over the range 45–55  C, and the extreme thermophiles (generally Archaea), some of which can grow at temperatures approaching 100  C. Most of the moderately thermophilic groups are neutrophiles. However, there are also neutrophiles such as Thermothrix (Tx.) thiopara and Sulfurihydrogenibium, which have a higher optimum growth temperature (72  C). These facultative autotrophs were found in neutral (pH 7.0), hot (74  C) springs (Caldwell et al. 1976; Brannan and Caldwell 1980; Takai et al. 2003). Thiothrix thiopara 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

Colorless Sulfur Bacteria

15

Molar ratio of inorganic sulfur compounds to organic substrates inorganic 100%

50:50

organic 100%

obligate chemolithotrophs

facultative chemolithotrophs

chemolithoheterotrophs

heterotrophs able to oxidize sulfur compounds

. Fig. 15.4 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 (seminatural) 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 15.4

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, the dissolved oxygen dropped from 3 mg 1 1 to 0.1 mg 1 1, but other parameters, such as pH and temperature, were unaffected. The Thiothrix 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 Archaea represent the colorless sulfur bacteria among the hyperthermophiles. 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). With growth temperatures between 60  C and 95  C, these species 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 15.4, 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, b). 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. 15.4). Thus, if the available substrate in energy-limited systems is wholly or predominantly inorganic, obligate autotrophs such as Halothiobacillus 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 P. 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 examples, see > Fig. 15.5a, b). Although mathematical modeling predicted that in some cases pure cultures of only one metabolic type should be obtained, in practice, satellite populations of the others remained (> Fig. 15.6). Clearly, secondary environmental or experimental conditions (e.g., excretion products such as glycolate, 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 determine the success of a particular species. Nevertheless, the use of this model (> Fig. 15.4) has now clarified the situation,

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a

b 100

100 Pa. versutus (+acetate)

90

PERCENTAGE OF TOTAL CELL-NUMBER

90

PERCENTAGE OF TOTAL CELL-NUMBER

566

80 70

Pa. versutus (+glycollate)

60 50 H. neapolitanus (+glycollate)

40 30 20 H. neapolitanus (+acetate)

10

Pa. versutus

80 70 60 50 40 30 20

spirillum G7

10

0

0 0

1

2

3

4

5

6

7

8

9

10

GLYCOLLATE OR ACETATE (mM)

0

1

2

3

4

5

6

7

8

9

10

THIOSULFATE (mM)

. Fig. 15.5 The effect of organic or inorganic energy sources on competition. (a) The effect of different concentrations of organic substrates on the competition between Paracoccus versutus (Pa. versutus) and Halothiobacillus neapolitanus (H. 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, Pa. versutus; closed symbols, H. neapolitanus; circles, glycolate supplied; triangles, acetate supplied. (b) The effect of thiosulfate on the competition for acetate (10 mM) between Paracoccus versutus (Pa. versutus) and a heterotrophic spirillum called G7. For experimental details, see (a). Open symbols, spirillum G7; closed symbols, Pa. versutus (Based on Gottschal et al. 1979)

a practical consequence being that it has shown the way for the selective enrichment of facultatively autotrophic sulfur bacteria from freshwater. 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. 15.5a, b, and 15.6), Gottschal et al. (1981) ran chemostat cultures alternating feeds of acetate and thiosulfate. In twomembered cultures, the mixotrophic P. 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 threemembered 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 re-induce its autotrophic enzymes each time, disappeared. As with the steady-state experiments, when different mixtures of acetate and thiosulfate alternated, the outcome was determined 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 before (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). Marine mixotrophs have been isolated, one of the earliest reports being a facultatively chemolithotrophic marine strain of Thiomonas intermedia from a thiosulfate-limited culture (Smith and Finazzo 1981). Similarly, an obligately autotrophic Thiomicrospira-type occurred in stable mixed cultures with a facultative Thioclava (Sorokin et al. 2005) when a facultative autotroph might have been expected. 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 pelophila and Thiobacillus thioparus. As can be seen from > Fig. 15.7, Thiomicrospira pelophila dominated mixed cultures of the two species at low iron concentrations, whereas Thiobacillus thioparus did better when iron was more abundant. One of the characteristics of Thiomicrospira pelophila is its tolerance of sulfide concentrations high enough to inhibit Thiobacillus

Colorless Sulfur Bacteria

100

15

µ 0.4

mixotroph

T. thioparus 0.3

80 percentage of total cell-number

Tms. pelophila 0.2

60

0.1

40

autotroph

heterotroph

20

Iron concentration

0 0

4

6 12 acetate (mM)

16

20

40

32

24 16 thiosulfate (mM)

8

0

1

2

. Fig. 15.7 The specific growth rates (m) of Thiomicrospira pelophila and Thiobacillus thioparus as a function of the iron concentration in chemostat cultures at 25 oC. 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)

3

. Fig. 15.6 Competition for acetate and thiosulfate in a chemostat between an autotroph, Halothiobacillus neapolitanus (open triangles); a mixotroph, Paracoccus 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 Pa. versutus dominated at intermediate ratios, as predicted, the other two types did not completely disappear. For the experimental details, see > Fig. 15.5a. 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. 1981; Gottschal and Thingstad 1982)

species. It is likely 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.

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. When Chester’s Manual of Determinative Bacteriology (a scanned copy of this forerunner of the famous Bergey’s Manual can now be downloaded from the Internet Archive; http://www.archive.org) appeared in 1901, it was based almost entirely on morphology, staining reactions, and growth on different media. Only two of the genera listed in > Table 15.2 were included – the morphologically distinctive Beggiatoa and

Thiothrix. Even today, morphology and the ability to grow on particular substrates, such as the reduced sulfur compounds, are still regarded as taxonomically significant. Needless to say, this has caused a certain amount of confusion (see > Table 15.2 for an overview of the genera involved and the most recent name changes). 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 consequently, 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 become 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 four distinct, if overlapping phases (morphology, physiology, analytical taxonomy and phylogeny), which will be discussed sequentially here.

Morphology The colorless sulfur bacteria include rods, spirals, cocci, filamentous cells, and it comes as no surprise to find that the first of

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Colorless Sulfur Bacteria

them to be described, Beggiatoa (Trevisan 1842), is also one of the largest. 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. As may be seen from a survey of taxonomic manuals, a few genera are still 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 ß-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 Paracoccus pantotrophus 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. 15.8a). Cultivation in batch culture on rich media in which rapid growth will occur leads to a slightly larger, pleomorphic form (> Fig. 15.8b). In chemostat cultures on mineral medium with acetate, chains of cocci appear. The internal structure of Paracoccus pantotrophus 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. 15.8c), gives way to cells with PHB granules and complex membranous structures (> Fig. 15.8d) 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. 15.8e), while denitrifying growth on sulfide can result in the accumulation of a fine deposit of sulfur in the periplasm (> Fig. 15.8f). The colonial form of this species also varies, with off-white, translucent colonies being produced during growth on mineral medium with acetate, hydrogen, or thiosulfate, and larger, thicker, browner colonies being generated during growth on rich media. Long-term continuous cultivation at high growth rates can select for faster-growing variants, giving rise to much bigger colonies when the culture is transferred to plates. Species with flagellae can lose them if shaken or stirred too quickly. 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 Halothiobacillus 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 well-defined conditions, this information should be used cautiously and in conjunction with other data.

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’ toolbox (Kluyver and van Niel 1936). For the obligate autotrophs, these might include such tests as optimum pH and 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 (Acidithiobacillus ferrooxidans and Thiobacillus 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 were late in 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 intended as an 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 apparently similar enzyme systems are. Thus, possession of the Calvin cycle enzymes for carbon dioxide fixation or the denitrification pathway enzymes is not considered sufficient grounds for 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, b) (see also > Fig. 15.2).

Analytical Taxonomy In many ways, the development of ‘‘analytical taxonomy’’ has been controlled by two factors – scientific knowledge and

Colorless Sulfur Bacteria

15

. Fig. 15.8 Variations in the morphology of cells of Paracoccus pantotrophus 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. (Figure b from Robertson and Kuenen 1983b. Figure c from Bonnet-Smits et al. 1988. Figure f, courtesy of H. J. Nanninga. All electron microscopy courtesy of W. Batenberg.) All bars = 0.5 mm

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. Table 15.5 Classification of species of colorless sulfur bacteria on the basis of their menaquinone and fatty acid composition Autotrophy type

Menaquinone

Hydroxy fatty acid

Species

Group

Facultative

MK-10

None

Starkeya novella

I.1

Facultative

MK-10

3OH 10:0

Paracoccus versutus

I.1

Facultative

MK-10

3OH 14:0

Acidiphilium acidophilum

I.2

Facultative

MK-8

3OH 10:0

‘‘Thiobacillus delicatus’’

II

Facultative

MK-8

3OH 10:0, 3OH 12:0

Thiomonas perometabolis

II

Facultative

MK-8

3OH 10:0, 3OH 12:0

Thiomonas intermedia

II

Facultative

MK-8

3OH 10:0, 3OH 12:0

Thiobacillus denitrificans

III.1

Facultative

MK-8

3OH 10:0, 3OH 12:0

Thiobacillus thioparus

III.1

Facultative

MK-8

3OH 12:0

Halothiobacillus neapolitanus

III.2

Facultative

MK-8

3OH 14:0

Acidithiobacillus ferrooxidans

III.3

Facultative

MK-8

3OH 14:0

Acidithiobacillus thiooxidans

III.3

MK menaquinone. The number indicates the number of isoprenoid units. Groupings are as proposed by Katayama-Fujimura et al. (1982)

available technology. Often, as an analytical technique became available, someone, somewhere, tested it for taxonomic significance. This section will provide a few examples. 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 can confirm that two strains are not related, matching %GC values do not guarantee that they are (see, for example, Kelly et al. 2005). Cellular fatty acid analysis has been used in the taxonomy of the Thiobacilli (Agate and Vishniac 1973). 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 15.5. 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 rRNA sequences (Lane et al. 1985; Stahl et al. 1987). This work was then extended by the use of 16S rRNA analysis (Lane et al. 1992) and revealed that there are closer matches between some sulfur-oxidizing bacteria and other apparently unrelated strains such as Escherichia coli than between these and other sulfur oxidizers. > Table 15.6 summarizes some of the results from the 5S and 16S rRNA comparisons. If the initial separation into obligate and facultative autotrophs employed by Katayama-Fujimura et al. (1982) is removed, the results shown in > Tables 15.5 and 15.6 apparently supported each other. Thus, groups I.1 and I.2 from the menaquinone/fatty acid analysis apparently correspond to group alpha from the 16S rRNA, groups II and III-1 with group beta-1, and groups III-2 and III-3 with beta-2. The

range of bacteria subjected to the menaquinone/fatty analysis was much smaller than that in the 5S and 16S rRNA survey. Comparison of the results in these tables with those showing the eventual outcome of the reorganization of the colorless sulfur bacteria shows some agreements among the obligate autotrophs. However, among the facultative species, the gulf between presumed related species (e.g., group 1.1. in > Table 15.5) has widened, and they are now believed to be separate genera. While chemotaxonomy and phylogeny might provide more reliable tools for the classification of these bacteria than physiological or morphological observations, the results are simply the best that can be obtained with current knowledge and technology. Microbial taxonomy has always been mutable. Phylogenic analyses of nucleic acids and proteins has revealed that physiological similarities are frequently coincidental rather than accurate indicators of relationships between microorganisms. Multilocus sequence analysis, rather than 16S rRNA analysis, has recently been gaining popularity. However, the pitfalls of relying on any single analytical system have been reviewed by Ka¨mpher and Glaeser (2012).

Phylogeny of Colorless Sulfur Bacteria Although novel opinions about defining prokaryotic species (Gevers et al. 2005) are numerous, 16S rRNA sequence analysis is still popular for the determination of the phylogenetic affiliation of species. Different databases giving rRNA sequences, such as the Ribosomal Database Project (RDP; Cole et al. 2009), the SILVA database (Pruesse et al. 2007), GreenGenes (DeSantis et al. 2006), EzTaxon (Chun et al. 2007), and the All-Species Living Tree Project (Yarza et al. 2010) are currently available. The principle of phylogenetic analysis (Felsenstein 2004) is simple, sequences are aligned to each other, and a phylogenetic tree is calculated using different algorithms (neighbor joining (Saitou and Nei 1988), maximum

Colorless Sulfur Bacteria

. Table 15.6 Classification of the colorless sulfur bacteria and examples of apparently related species (group ‘‘purple’’), also termed Proteobacteria (Stackebrandt et al. 1988), as shown by 5S and partial 16S rRNA analysisa Main group Subgroup Species a

b

1

Acidiphilium acidophilus, Acidiphilium rubrum

1

Acidiphillium cryptum, Starkeya novella

2

Rhodobacter capsulatus, Paracoccus versutus

2

Paracoccus denitrificans

1

Thiobacillus denitrificans, Thiobacillus thioparus

1

Thiomonas intermedia, Thiomonas perometabolis

1

Rhodocyclus gelatinosus

1

Vitreoscilla

Borderline Halothiobacillus neapolitanus, Chromatium vinosum g

gb

Delta

1

Thiothrix nivea, Riftia symbionts

1

Thiomicrospira pelophila, Thiomicrospira L-12

1

Bathymodiolus symbionts

1

Other symbionts

1

Pseudomonas aeruginosa, Pseudomonas putida

1

Beggiatoa alba, Beggiatoa sp.

2

Escherichia coli, Salmonella, Proteus, Vibrio

3

Thermithiobacillus tepidarius, Acidithiobacillus ferrooxidans

3

Acidithiobacillus albertensis, Acidithiobacillus thiooxidans Thiovulum, Campylobacter, Wolinella

Atypical strains have been omitted for the sake of simplicity a Adapted from Lane et al. 1992 and Harrison 1989 b The organisms shown here as ‘‘g3" were originally shown as being on a ß side branch, but improved techniques have now shown them to be in group g (Kelly et al. 2000)

parsimony, maximum likelihood (Saitou and Nei 1988) and evolutionary models (e.g., Kimura 1980). Special programs, such as ARB (Ludwich et al. 2004), are available for this purpose, but different tools to create phylogenetic trees are also incorporated in some of the rRNA databases mentioned above. Based on comparative analysis of 16S rRNA sequences, the known colorless sulfur bacteria are, at the time of writing, grouped into four phylogenetic lineages, three within the Bacteria and one within the Archaea (> Fig. 15.9). Most of the colorless sulfur bacteria belong to the phylum Proteobacteria, in particular the class Gammaproteobacteria. The group named

15

Thiomicrospira also includes four Thioalkalimicrobium species (Thioalkalimicrobium aerophilum, Thioalkalimicrobium microaerophilum, Thioalkalimicrobium sibiricum, and Thioalkalimicrobium cyclicum), which are closely related to Thiomicrospira pelophila and Thiomicrospira thyasirae. Other groups of colorless sulfur bacteria within the Gammaproteobacteria are Thiothrix (nine species), Halothiobacillus (four species), Thiohalomonas (two species), Thiohalospira (two species), Thioalkalivibrio (nine species), and the Acidithiobacillaceae (five species). Candidatus Thiomargarita namibiensis, which has not been isolated in pure culture, is related to Thioploca ingrica and Beggiatoa alba. Only a few of the colorless sulfur bacteria belong to the other classes within the Proteobacteria; five Thiobacillus spp. (Thiobacillus denitrificans, Thiobacillus thioparus, Thiobacillus thiophilus, Thiobacillus aquaesulis, and Thiobacillus subterraneus) as well as Sulfuricella denitrificans (Kojima and Fukui 2010) are grouped within the Betaproteobacteria. Starkeya koreensis, Starkeya novella, and Thioclava pacifica (Sorokin et al. 2005) belong to the Alphaproteobacteria, and Sulfurimonas autotrophica, Sulfurimonas denitrificans, Sulfurimonas paralvinella, and Sulfurovum lithotrophicum, grouped as Sulfurimonas, are belonging together with Thiovulum to the Epsilonproteobacteria. So far, there are no colorless sulfur bacteria known that belong to the Deltaproteobacteria. In addition to the Proteobacteria, five species belong to the genus Sulfobacillus within the phylum Firmicutes, and five others are belonging to the genus Sulfurihydrogenibium within the phylum Aquificae. Within the Archaea, six Sulfolobus species, four Acidianus species, and Sulfurisphaera ohwakuensis belong to the family Sulfolobaceae within the phylum Crenarchaeota. Comparative 16S rRNA sequence analysis is not only commonly used to determine the phylogenetic position of novel species, it is also used to reclassify the phylogenetic position of existing species or to solve different taxonomic problems. Recently, Boden et al. (2011) performed a phylogenetic assessment of 12 strains of Thiobacillus thioparus present in different culture collections in order to check whether the so-called Starkey type strain deposited in the different collections was identical, to obtain a definitive reference 16S rRNA sequence, and to check of other strains labeled as Thiobacillus thioparus were well-founded examples of the species. They found that four examples of the Starkey type strain were identical, and so they could obtain a definitive reference sequence. Comparative sequence analysis subsequently showed that 6 strains were correctly affiliated to Thiobacillus thioparus, but that two strains were wrongly named and had to be renamed as Halothiobacillus neapolitanus and Thermithiobacillus tepidarius, respectively. Apart from the use of 16S rRNA sequences, the phylogeny of sulfur-oxidizing bacteria, including colorless sulfur bacteria, has also been studied using the soxB gene (Meyer et al. 2007), the aprBA genes (Meyer and Kuever 2007b), and RuBisCO (Tourova et al. 2006). Although in general the phylogenies based on these genes are similar to the ones inferred from 16S rRNA sequence data, there might be differences due to lateral gene transfer (see Meyer and Kuever 2007a).

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Colorless Sulfur Bacteria

. Fig. 15.9 Phylogenetic tree based on nearly complete 16S ribosomal RNA (rRNA) sequences of described colorless sulfur bacteria. The sequences were obtained from ‘‘The All-Species Living Tree’’ project (LTP release 106; Yarza et al. 2010), which contains high-quality, curated 16S rRNA sequences of type strains. The tree was created using ARB software. The number within the collapsed clusters indicates the number of different species within a particular group. The scale bar indicates 10 % sequence difference

Molecular Methods to Study the Diversity of Colorless Sulfur Bacteria in Natural Habitats Detection of Colorless Sulfur Bacteria Using Molecular Markers It is now well recognized that less than 1 % of all bacteria in nature can be isolated in pure cultures. In order to study the diversity and activity of microbial communities and the dynamics of their community members, it is necessary to use molecular techniques (e.g., polymerase chain reaction (PCR), denaturing

gradient gel electrophoresis (DGGE), fluorescence in situ hybridization (FISH), and next-generation sequencing (NGS)) that were originally developed in molecular biology and medicine. Thus far, most of the bacteria, including the colorless sulfur bacteria, have been detected and identified by using the 16S rRNA approach where DNA fragments PCR-amplified with primers targeting the 16S rRNA gene of Bacteria or Archaea were sequenced after cloning or DGGE analysis. By using this approach, Hubert et al. (2011) could show a massive dominance of sequences (87 %) closely related to the chemolithoautotrophic sulfur-oxidizing bacterium Sulfuricurvum kujiense in formation

Colorless Sulfur Bacteria

waters from a Canadian oil sand reservoir. Lanse´n et al. (2011) used the 16S rRNA approach to detect members belonging to the epsilonproteobacterial genera Sulfurimonas and Sulfurovum in microbial communities of the Jan Mayen hydrothermal vent field. Yang et al. (2011) detected the dominance of members affiliated to the genus Sulfurihydrogenibium (Aquificales) in hydrothermal vents of Yellowstone Lake with temperatures of 50–60  C, while members related to the gammaproteobacterium sulfur oxidizer Thiovirga were more dominant in vents with lower temperatures. To quantify the number of cells and to study their spatial distribution, fluorescence in situ hybridization (FISH) is used. The principle of FISH is straightforward. Bacterial cells are fixed in paraformaldehyde and dehydrated in ethanol. Subsequently, the cells are hybridized with oligonucleotide probes labeled with different fluorescent dyes and visualized with an epifluorescence microscope (Amann and Fuchs 2008). Maestre et al. (2010) used 16S rRNA gene libraries and FISH to study the bacterial community in a laboratory-scale biotrickling filter treating high loads of H2S and found that most of the community members were affiliated to Thiothrix, Thiobacillus, and Sulfurimonas denitrificans. A similar approach was used by Okabe and coworkers (2007) to follow the succession of sulfur-oxidizing bacteria (SOxB) in microbial communities involved in concrete corrosion of sewer systems. They found that six different phylotypes of SOxB were present and that their abundance shifted in the following order: Thiothrix sp., Thiobacillus plumbophilus, Thiomonas intermedia, Halothiobacillus neapolitanus, Acidiphilium acidophilum, and Acidithiobacillus thiooxidans. Another method for quantifying the number of bacteria is quantitative PCR (qPCR). Reigstad et al. (2011) used qPCR of 16S rRNA genes to study microbial communities in thermal springs on Svalbard and found that bacteria closely related to Thiothrix and Sulfurovum were the most dominant constituents of these communities. Liu et al. (2006) designed specific primers targeting the 16S rRNA genes of different microorganisms involved in bioleaching, including the colorless sulfur bacteria Sulfolobus, Sulfobacillus, and Acidithiobacillus caldus, and used these primers in a qPCR to rapidly detect and quantify these organisms in bioleaching processes. A combination of different molecular methods (qPCR, FISH, CARD-FISH) and most probable number (MPN) cultivation techniques was used by Kock and Schippers (2008) to quantitatively analyze microbial communities from three different sulfidic mine waste tailing dumps. They found that Acidithiobacillus spp. dominated over Leptospirillum spp. and that Sulfobacillus spp. were generally less abundant. Apart from the use of 16S rRNA genes as molecular markers, functional genes were also used to study colorless sulfur bacteria in their natural habitat. Meyer and Kuever (2007b) used aprA as a functional marker to study the diversity of sulfate-reducing and sulfur-oxidizing prokaryotes in different samples from the Caribbean Sea and found a dominance of putative chemolithoheterotrophic sulfur-oxidizing Alphaproteobacteria in nonhydrothermal sediments and in the water column and

15

chemolithoautotrophic sulfur-oxidizing Beta- and Gammaproteobacteria on the surface of volcanic manganese crusts. Sorokin and coworkers used RuBisCO and ATP lyase genes to study the diversity of sulfur-oxidizing bacteria in hypersaline (Tourova et al. 2010) and haloalkaline (Kovaleva et al. 2011) lakes. Chen and coworkers (2007) used both 16S rRNA and sulfur oxygenase reductase (SOR) genes as molecular markers to study the microorganisms in bioreactors treating goldbearing concentrates.

Natural Habitats of Colorless Sulfur Bacteria 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. Indeed, where they are very active, the reduced sulfur compounds may not reach detectable levels. Because the range of habitats is so wide, the principles underlying the selection of colorless sulfur bacteria in selected situations will be discussed below. The following section will then deal more generally with the role of the colorless sulfur-oxidizing bacteria in the sulfur cycle. 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 sulfides (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 sulfidecontaining areas and aerobic waters and sediments. There, 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, and the acidophilic bacteria need not, therefore, grow 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 at any pH. Another habitat in which sulfideoxidizing 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. As well as free-living species such as Thiomicrospira crunogena (Jannasch 1985), it has been 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.

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Colorless Sulfur Bacteria

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 hundred micrometers 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 sp. 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 seawater 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 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 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). 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. The turnover time for sulfide and oxygen was only 3 s 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 clear-cut. Schmidt et al. (1987) showed sulfide oxidation rates for a freshwater strain comparable to those obtained with the marine strain discussed above. 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 1 mM 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 turnover of inorganic and organic compounds is heavily dominated by the sulfur cycle) on the surface of sediment. It was observed that the mixture of cyanobacteria, phototrophic sulfur bacteria, and Beggiatoa was stratified and that the relative positions of the three populations among the strata were governed by the level of photosynthetically generated oxygen (> Fig. 15.10). 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 Achromatium 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 sulfide-dependent 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 (see Jannasch 1985, 1988 for early reviews). 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 seawater, forming precipitates that often accumulate as ‘‘chimneys.’’

Colorless Sulfur Bacteria

distance/mm

−1

15

O2

0 Oscillatoria 1 2

Beggiatoa Chromatium

H2S 12

distance/mm

−1

09

O2

15 day

0

O2

Beggiatoa 1

06 H2S

18

Oscillatoria

Beggiatoa night

Chromatium

Oscillatoria

2 03

21

H2S

Chromatium

24

distance/mm

−1

O2

Chromatium

0 Beggiatoa 1

H2S

2

Oscillatoria Chromatium

. Fig. 15.10 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)

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 ( ‘‘Symbiosis’’ below). In addition to sulfide concentration and temperature, the authors suggested that organic excretion compounds from the worms may be an important factor in the development of these mats. Beggiatoa mats are also strongly affected by the patchy nature of hydrothermal seeps (Lloyd et al. 2010) and the rate of pore water flow (de Beer et al. 2006) since these control the supply rates for electron donors and acceptors to the mat.

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Symbiosis Geologists studying areas of volcanic activity on the seabed (1,800–3,700 m below the surface) were surprised to find that not only were there dense, free-living bacterial populations associated with the vents, but that these permanently dark areas were also occupied by an extensive community of invertebrates (and also, in some areas, fish), most of which were previously unknown (Corliss et al. 1979). Despite the density of the bacterial community, it was difficult to see how a food chain based entirely on suspended bacteria as a source of prey could support the considerable population of very large tube worms, clams, and other invertebrates. Investigation of the anatomy of the tube worms (Riftia pachyptila) revealed that they do not have an alimentary tract but instead possess a large (more than half the weight of the worm) body of tissue, the trophosome, which is very rich in blood vessels. Examination of this tissue under the electron microscope revealed that it also contained a dense, intracellular community of bacteria (Cavanaugh et al. 1981; Cavanaugh 1983a). The trophosome had already been shown to contain the enzymes necessary for chemoautotrophic growth on reduced sulfur compounds. These enzymes did not occur elsewhere in the tissues of the worm (Felbeck 1981; Felbeck et al. 1981) and were presumably derived from the bacteria. This was the first example of prokaryotic/eukaryotic symbiosis in which the hosts rely on organic compounds excreted by the bacteria. The blood of the tube worms carries sulfide as well as oxygen from the gills to the trophosome and has a special sulfidebinding protein that prevents sulfide toxicity. Endosymbionts were then found in a range of vent faunas including the giant white clams (Calyptogena magnifica) and were not limited to sulfide-oxidizing bacteria, since methylotrophs have also been found (Jannasch 1988). 5S and 16S rRNA analysis indicated a relatively close relationship between the endosymbionts and members of the genus Thiomicrospira (Lane et al. 1985, 1992), which, as mentioned above, is one of the best-represented genera among the free-living bacterial community at the vents (Ruby et al. 1981; Ruby and Jannasch 1982; Jannasch 1988). Once the occurrence of endosymbiotic bacteria in the animals of the hydrothermal vents had been accepted, many more occurrences were recognized in more mundane locations, including sewage outfalls and sulfide-rich sediments (e.g., Southward 1986; Dando and Southward 1986). Many of the animals associated with symbionts resemble Riftia pachyptila in that they completely lack a mouth and digestive system, whereas others may have only small guts and feeding appendages (Cavanaugh 1983a, b). Not all of them have a specialized organ like the trophosome, and many endosymbionts appear to be associated with the gills of the eukaryotic host. For example, intracellular colorless sulfur bacteria have been found in the gill tissues of bivalves such as Solemya velum (Cavanaugh 1983b) and Thyasira flexuosa (Wood and Kelly 1989). The description of a novel Thiobacillus-like species, Thiomicrospira thyasiridae (Wood and Kelly 1989), from the gill tissue of Thyasira flexuosa was probably the first report of the isolation of one of these symbionts.

Smith et al. (1989) illustrated the effect that a localized deposit of organic material in an otherwise oligotrophic environment can have on the indigenous community. The skeleton of a 20-m-long whale at a depth of 1,240 m on the seabed in the Santa Catalina basin was not only covered with mats of Beggiatoa resembling Beggiatoa gigantea, but it also supported six metazoan species, at least four of which are known in other locations to contain endosymbionts. As well as vent species (Vesicomya gigas and Calyptogena pacifica), others organisms known from anoxic sediments (Lucinoma annulata) and rotting wood (Idasola washingtonia) were also observed. None of these prokaryotic or eukaryotic species had been observed in this area before. It was found that the pore water under the skeleton contained around 20 mM sulfide, and the samples of whalebone that were recovered were found to be rich in oil and smelled strongly of sulfide. It would appear from the apparent ages of some of the mollusks present that a single whale carcass is sufficient to support these sulfide-dependent communities for several years.

Man-made Habitats and Application of Sulfur Bacteria Man-made environments such as the bioreactors used for industrial wastewater treatment have provided habitats for bacteria that impose selective parameters not necessarily found in nature. Thus, substrates tend to be more abundant and conditions are generally more stable than in most natural situations. Two categories of artificial habitat where colorless sulfur bacteria are particularly important are wastewater treatment bioreactors and those associated with various leaching activities. Examples of other artificial habitats include industrial sulfur deposits or dumps, mining operations that expose sulfidic ores or sulfur to water or air, coal storage sites, and, last but not least, systems (including sewage treatment plants) containing various amounts of reduced sulfur compounds.

Waste Treatment Reduced sulfur compounds can occur in industrial wastes in a variety of forms and from a variety of sources. For example, sulfide is an inevitable by-product of sulfate reduction associated with methanogenesis (if the effluent from which the methane is being generated contains significant amounts of sulfate) and the oil and gas industries. Thiosulfate and thiocyanate make up a substantial amount of the chemical content of photographic film processing waste, and some papermaking processes generate both inorganic and organic sulfides. Of course, the amount of reduced sulfur compounds generated from industrial processes pales into insignificance when the quantity generated from animal wastes is considered. Reduced sulfur compounds present a problem both environmentally, because of their toxicity, and socially, because of their odor. If large amounts of sulfide are released into natural

Colorless Sulfur Bacteria

waters, this can result in oxygen depletion, either because of the oxygen demand for biological oxidation or, in the absence of suitable bacteria, by spontaneous chemical oxidation. Many water treatment plants impose surcharges for the treatment of such effluent because it can disturb the microbial community in the bioreactors, and there is obviously considerable pressure on companies to treat their effluent on the site. There are both chemical and physical methods of removing hydrogen sulfide from effluent; these include the use of ion-exchange resins, absorption with aqueous or organic solvents, and chemical oxidation (Gommers 1988). Many of these simply transfer the problem to another waste stream or involve expensive or complex processes, and they are all expensive, especially for the removal of the last traces of sulfide compounds. Colorless sulfur bacteria occur in many sewage treatment systems and, in fact, are inadvertently used to oxidize reduced sulfur compounds in the wastewater. In some cases, this can lead to problems, such as the ‘‘bulking’’ caused by Thiothrix. The deliberate use of biological treatment of sulfide-containing waste using colorless sulfur bacteria has become commonplace. The end products (sulfur or sulfate) are not hazardous, and sulfate can be discharged directly into the sea or into brackish estuaries (which already are so high in sulfate that the discharge is insignificant). Moreover, biological treatment systems can be based on existing reactor designs (e.g., fluidized and packed bed reactors) and require very little in the way of new technology. Another advantage of a biological process is that it can be combined with the treatment of other problems in an effluent. For example, the effluent of a methane reactor will contain ammonia in addition to sulfide. If the ammonia is then converted to nitrate or nitrite by aerobic, nitrifying bacteria, the resulting effluent can then be recycled to provide the electron acceptor for a sulfide-oxidizing reactor immediately after the methane reactor. The microbiological investigation of such a sulfide-oxidizing, denitrifying reactor revealed the presence of large numbers of facultatively autotrophic colorless sulfur bacteria, which could oxidize sulfide to sulfate while reducing nitrate to nitrogen gas (Robertson and Kuenen 1983a). In addition to the removal of nitrogen compounds, other advantages associated with the use of denitrifying bacteria rather than aerobic ones include lower production of both biomass and acid.

Combined Sulfide Oxidation and Denitrification A denitrifying, sulfide -oxidizing reactor system was patented by a Dutch company, Gist-brocades (now a branch of DSM), for the posttreatment of effluent from methane-producing reactors (Patent number E.P.A.0051 888). Studies on a laboratory-scale model of this reactor, running on artificial wastewater, revealed that sulfide (2–3 kg S/m3/day), acetate (4–6 kg S/m3/day), and nitrate (5 kg S/m3/day) were all effectively removed (Gommers et al. 1988a). The rate-limiting step in the reactor proved to be the oxidation of sulfur to sulfate and, under most loads, the biomass had an overcapacity for both the oxidation of sulfide to sulfur and the conversion of acetate (Gommers et al. 1988b).

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During experiments in which nitrate depletion occurred, it became evident that in the absence of nitrate, at least one member of the bacterial community was able to reduce any available sulfur, thus illustrating the need for careful monitoring of the electron donor/electron acceptor ratios in such reactors (Gommers et al. 1988b). The facultatively autotrophic species Paracoccus pantotrophus was isolated from a denitrifying, sulfide-oxidizing uidized bed reactor that was supplied with approximately equivalent amounts of organic and inorganic substrates (Robertson and Kuenen 1983b), and it initially appeared that the selection of a facultative bacterium would lend support to the model described for the ecological niches of aerobic, freshwater sulfur-oxidizing bacteria (> Fig. 15.4). However, subsequent attempts to isolate obligate autotrophs from a laboratory-scale model of this system that was being fed with an exclusively inorganic feed also resulted in the isolation of facultative autotrophs (M. Verbeek, W. Bijleveld, L. A. Robertson, and J. G. Kuenen, unpublished observations). It is not clear whether obligate autotrophs were present in the inoculum, or the isolation techniques employed were inadequate for any obligate autotrophs present (although they were adequate for the cultivation of known obligate autotrophs), or whether growth in a biofilm in this type of reactor poses an additional selective pressure that favors facultatively autotrophic bacteria. Subsequent work has shown that a number of sulfide oxidizers from a wastewater system required cultivation on special membrane filters with sulfide gas before isolated colonies could be obtained (Visser et al. 1997). The same basic idea of using denitrifying colorless sulfur bacteria was employed in a method proposed by Sublette and Sylvester (1987) for removing H2S from gas streams by passing them through a reactor containing Thiobacillus denitrificans. The bacteria were first immobilized by coculturing with floc-forming heterotrophs after the authors demonstrated that the presence of the heterotroph had no effect on the sulfide oxidation rate of Thiobacillus denitrificans.

Removal of Sulfide as Elemental Sulfur As already mentioned, sulfate-containing effluents can be discharged into the sea without significantly increasing the sulfur budget. However, the same is not true if the effluent is discharged into a body of freshwater. To overcome this problem, recovery as elemental sulfur, an intermediate in the oxidation of sulfide to sulfate, would be more appropriate. Research has shown that certain Thiobacillus-like bacteria are more inclined to produce sulfur than other species and that both the dissolved oxygen and the sulfide concentration play an important part in determining whether sulfur or sulfate is the primary end product during sulfide oxidation. Both electron acceptor limitation and high sulfide loads favor sulfur production (Stefess and Kuenen 1989). A pilot plant based on this principle, using a mixed bacterial biofilm reactor to treat the effluent from a paper mill, was developed in the Netherlands (Buisman 1989; Janssen et al 2009).

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Removal of Organic Sulfides A problem frequently encountered during the alkaline pulping of wood is the production of organic sulfides such as methyl mercaptan and dimethyl sulfide. Alkaline pulping is done in order to improve the yield and quality of pulp derived from conifers to be used primarily in the manufacture of paper. Organic sulfides are toxic at even lower concentrations than hydrogen sulfide and have a very low threshold odor. Despite their toxicity, it has proved possible to grow bacteria on high concentrations of organic sulfides by using substrate-limited chemostats (Suylen et al. 1986; Kanagawa and Kelly 1986; Smith and Kelly 1988a, b, c). That the ability to oxidize these compounds may be widespread is suggested by the observation that the dominant organism in one set of experiments was a Hyphomicrobium species that was later shown to be able to grow as a facultative chemolithotroph on organic sulfur compounds in pure culture (Suylen and Kuenen 1986; Suylen et al. 1986), whereas the key organism in the other series was a strain of Thiobacillus thioparus, an obligate autotroph (Kanagawa and Kelly 1986). Immobilized cells of Thiobacillus thioparus strain TK-m have been successfully used on the laboratory scale to deodorize gases containing methyl mercaptan, dimethyl sulfide, dimethyl disulfide, and hydrogen sulfide (Kanagawa and Mikami 1989; Tanji et al. 1989). All of the colorless sulfur bacteria mentioned thus far are beneficial in wastewater treatment. However, in oxidation tanks fed with sulfide-containing waste water, the filamentous Thiothrix species can cause problems because they are associated with the phenomenon known as ‘‘bulking’’; this occurs when bacterial aggregations that usually settle easily become loose and flocculent. This can result in blockages or loss of the biomass from the reactor.

Leaching-Associated Activities Acidophilic bacteria are used in the recovery of metals from poor ores by leaching, and their potential use in the desulfurization of coal was extensively studied in the later decades of the twentieth century. To some extent, coal desulfurization and microbial leaching are the same process, in that in both cases sulfidic ores are oxidized, using similar organisms. However, the desired end products are different, and they are thus generally discussed separately. The aim of coal desulfurization is to produce a solid product (coal) that is as free of sulfur (including sulfur-containing precipitates) as possible, and it is therefore necessary to convert reduced sulfur compounds to soluble forms. In leaching, it is metal recovery that is important, and the presence of jarosite (M∗Fe3(SO4)2OH5, where M is a monovalent cation such as Na+ or K+) and other precipitates in the solid waste is not relevant (although it may constitute an environmental problem around the leaching heaps).

Bacterial leaching is used in the recovery of metals from ores that are too poor for conventional metallurgical extraction methods (Kelly and Tuovinen 1972; Brierley and Lockwood 1977; Brierley 1982; Ehrlich and Brierley (1990)). Combinations of Acidithiobacillus ferrooxidans and either A. thiooxidans or Acidiphilium acidophilum and Leptospirillum ferrooxidans have been associated with the degradation of pyrite (FeS2) and chalcopyrite (CuFeS2). The leaching reactions may involve the direct bacterial oxidation of the sulfide ores with oxygen and/or an indirect process during which ferric ions produced by the bacterial oxidation of ferrous iron are used to chemically oxidize the sulfide ores. The ferric ions are thereby reduced to ferrous iron, which, in turn, can be recycled by the bacteria. During this process, other metallic ions such as cupric copper dissolve. Other metals that have been extracted using processes that involve bacteria include zinc, uranium, lead, gold, molybdenum, and, especially, copper. Dump leaching operations, which are frequently used to extract copper, can be fairly primitive, involving the creation of ore dumps, often in valleys or old open pit mines. As water percolates through the heaped rocks, bacterial activity releases the metals into solution. This solution is then collected in catch basins, the metals recovered, and the liquids recycled to the top of the dump. A somewhat better controlled system is known as heap leaching. During this process, the ore-bearing rocks are crushed to promote contact with the acidified water, and the heaps are built on impermeable bases that prevent seepage into the soil beneath. Aeration systems can be built into the heaps. As mineral reserves become depleted and demand increases, it is becoming economically attractive to extract even small amounts of metals in poor ores and spoilage heaps, technological improvements should increase the efficiency of microbial leaching processes and lessen their environmental impact. Bioleaching of other potential sources such as mine tailings, contaminated sediments, and even sewage sludge is also attracting attention (Liu et al. 2008; Seidel et al. 2006; Pathak et al. 2009). Research into the use of the pyrite-oxidizing abilities of bacteria such as Acidithiobacillus ferrooxidans and Sulfolobus species for the removal of sulfur compounds from coal before it is burned, thus reducing sulfur emission into the atmosphere, has been carried out at a number of centers in the last few decades. It has been shown that such a process could be effective, especially for low-sulfur coals, using consortia of mesophilic bacteria (Bos et al. 1988; Bos and Kuenen 1990). Laboratory studies have shown that an optimal process requires two steps. First, a mixed-flow inoculation step, where a fairly dense population of bacteria already growing on pyrite can be brought into contact with fresh, finely ground coal at a pH suitable for growth (around pH 1.8). This inoculation step would then be followed by the use of plug-flow reactors, where the bulk of the pyrite oxidation would take place. At the end of the process, the process water can be recirculated, as can some of the biomass-bearing coal particles, to serve as the inoculum for the fresh coal. A plant design, involving a cascade of Pachuca tanks (> Fig. 15.11), was devised for this type of system (Bos et al. 1988). Pachuca tanks

Colorless Sulfur Bacteria coal

the water, but also with the toxic concentrations of heavy metals that they may contain. In addition, acidic water containing ferric sulfate may generate precipitates of jarosite, and these can block drainage pipes and cover stream and river sediments.

grinding

nutrients bio-desulphurisation reactors

water

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washing

compressor

dewatering Fe and SO4 removal

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drying desulphurised coal

waste

. Fig. 15.11 Simplified scheme for the microbiological desulfurization of coal in Pachuca tanks (biodesulfurization reactors). After grinding and mixing with water, the coal slurry contains particles less than 100 mm in diameter at a concentration of 20 % (w/v). At this particle size, virtually all of the pyrite crystals become accessible to microbial leaching. The total leaching process requires about 10 days. More than 95 % of the inorganic sulfur is removed, but little or no organic sulfur is degraded

(in their simplest form, an inverted cone with aeration at the narrowest point, at the bottom of the tank) are particularly suitable for this type of process because the upflow of air into the tanks not only provides the bacterial community with the oxygen and carbon dioxide necessary, but it also keeps the slurry well-mixed, without any need for complex and expensive stirring mechanisms.

Corrosion Together with the sulfate-reducing bacteria, many of the sulfuric-acid-producing bacteria, and in particular the acidophiles, have been implicated in many corrosion problems. Indeed a strain of Acidithiobacillus thiooxidans isolated from a corroding concrete pipe was originally known as Thiobacillus concretivorus (i.e., concrete-eating). In sewage pipes with an aerobic headspace, sulfide may be produced in the anaerobic water phase and then be transferred to the film of water on the aerobic part of the pipe where it may be oxidized to sulfuric acid. In order to dissolve the carbonates in concrete, the pH need only be below 5.0–5.5 and such a pH can be generated by either neutrophilic or acidophilic bacteria. The activities of the acidophiles may also be responsible for steel pipe corrosion (Kuenen and Bos 1988) as well as many of the pollution problems associated with the acid runoff from mine spoil heaps. These environmental problems are not only associated with the low pH of

The Role of Colorless Sulfur-Oxidizing Bacteria in the Sulfur Cycle Although much is known about the physiology and occurrence of colorless sulfur bacteria (especially with the advent of gene detection and DNA analysis methods), less is known about the quantitative aspects of their activity in nature. Many of the reasons for this are difficulties commonly associated with field work (e.g., heterogeneous samples, unstable gradients, low concentrations of substrates) and are therefore outside the scope of this chapter, but a few difficulties are uniquely associated with the colorless sulfur bacteria. Commonly used methods for estimating the activity of sulfur-oxidizing bacteria in the field include cell counts, oxidation of (radiolabeled) substrate (sulfide, thiosulfate, or sulfur), product formation (especially sulfuric acid, since this causes pH changes), and 14CO2 fixation. Other, more specific techniques include the measurement of substrate-dependent respiration and immunofluorescent microscopy.

Cell Counts With some of the more conspicuous bacteria (e.g., Beggiatoa, Thiovulum), it is possible to obtain a rough estimate of numbers based on direct cell counts. However, most of the colorless sulfur bacteria require cultivation before they can be counted. The choice of media and substrates for most probable number (MPN) estimates or direct plate counts is especially difficult for the colorless sulfur bacteria. The most obvious problem is that outside the chemostat there is no way of selectively growing facultative autotrophs or chemolithoheterotrophs. They must first be isolated on autotrophic or heterotrophic media, respectively, and then screened for sulfur-oxidizing capacity. In addition, low recovery efficiency can be a problem with both plate counts and dilution series. Two other problems are associated with the obligate autotrophs. Firstly, thiosulfate is frequently used as an energy source in solid media, but this is not always the most suitable energy source. For some bacteria, agar plates containing colloidal sulfur may be more appropriate, while other bacteria may require sulfide. The use of solid sulfide media can present technical problems with regard to toxicity and instability unless one of the less-soluble nontoxic sulfides (e.g., calcium sulfide) is used. Secondly, some autotrophic species do not give distinct colonies on agar, and moreover, the acidophiles may be inhibited by organic compounds resulting from chemical acid hydrolysis of the agar itself at their required growth pH values. To overcome these agar-associated problems, other techniques, such as the use

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of silica gel plates or floating filters (de Bruyn et al. 1990; Visser et al 1997), may be more appropriate. Some of the sulfuroxidizers may have a requirement for an unidentified growth factor such as a vitamin or mineral.

1

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Activity Measurements Data on the rates of sulfide oxidation in natural systems are scattered and somewhat variable, possibly because of the difficulty of accurate sampling as well as the reactivity of the compounds involved. Once cell numbers have been estimated with a degree of confidence, they can only be used to provide an idea of the potential activity of colorless sulfur-oxidizing bacteria within that particular ecosystem. The measurement of substrate transformations (i.e., utilization or accumulation), preferably in situ, can be used as a measure of actual activity. A major problem associated with the use and measurement of many reduced sulfur compounds, especially sulfide and sulfite, is that they are chemically very reactive and are readily oxidized spontaneously by oxygen. Appropriate controls can, to some extent, overcome this problem, but it must be remembered that in nature biological and chemical reactions compete, and equilibrium reactions causing the exchange of radiolabel in reduced sulfur compounds mean that extra caution must be used in the interpretation of results. Moreover, chemical oxidation rates are influenced by many of the environmental parameters that also affect biological activity (e.g., pH, temperature, chemical constitution of the solutions involved). In a few cases, where dominant populations of known colorless sulfur bacteria occur (e.g., Sulfolobus in solfataras, Beggiatoa mats), rough estimates have been made of the activity of these organisms. Mosser et al. (1973) found rates for sulfur oxidation to sulfate of 67 and 190 g m 2day 1 for mats of Sulfolobus acidocaldarius growing in two hot pools (Moose Pool and Sulfur Cauldron, Yellowstone National Park, respectively). In the Black Sea, a maximum rate of 710 nmol l 1day 1 was observed by Sorokin (1970). For extended discussions of sulfur oxidation rates in nature, the reader is referred to Kuenen (1975) and to Jørgensen (1988). Another problem is that the sulfur-oxidizing heterotrophs may also contribute to the turnover of reduced sulfur compounds at natural sites. In some cases, 14CO2 fixation can be used to eliminate this, but in many locations where mixotrophs or chemolithoheterotrophs are involved, CO2 may not be the primary source of carbon. This type of experiment could, therefore, sometimes result in underestimates if it is not used in tandem with other measurements. An associated problem is that the specific activity of a given species can vary. For example, Beudeker et al. (1980) found that, when grown under carbon dioxide limitation, the ribulose bisphosphate carboxylase (RuBisCO) activity in Halothiobacillus neapolitanus was 240 nmol min 1mg protein 1. If, however, thiosulfate was the limiting factor, the enzyme level fell to 72 nmol min 1mg protein 1. Other substrate conversion rates can also vary, especially among species. Thus, it has been found that Thiobacillus denitrificans

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. Fig. 15.12 Profiles of CO2 fixation, dissolved oxygen, and dissolved hydrogen sulfide concentrations in Saelenvaan Lake (Norway) sampled at 5 a.m. on 15 August 1978. The high resolution was due to the use of a special sampling device connected to a pump. CO2 fixation rates (top horizontal axis) were obtained using 14CO2 injected into dark bottles, which were incubated in situ. The left vertical axis is depth in m. The bottom horizontal axis is the concentration of dissolved gas in mmol l 2. Triangles, mmol CO2 l Eh 1; squares, mmol oxygen liter1; circles, mmol hydrogen sulfide liter–1 (From Kelly and Kuenen 1984)

and Sulfurimonas denitrificans oxidize thiosulfate at rates of 0.86 and 2.9 mM thiosulfate g C 1h 1, respectively (Timmer ten Hoor 1977). A combination of CO2 fixation and oxygen and hydrogen sulfide analysis was used to measure microbial activity in Saelenvaan Lake in Norway. As can be seen from > Fig. 15.12, a peak of CO2 fixation was found to coincide with the very narrow zone where oxygen and sulfide coexisted. It should be noted that the sampling technique was critical for the success of these experiments. A special sampling device with an inlet that removes water from a horizontal area of the column at 1–2 cm intervals (Jørgensen et al. 1979) was necessary if a less accurate device was used, the very narrow CO2 fixation zone could not be seen because of dilution by the surrounding water. Those working on the ecosystems around the hydrothermal vents have, of course, severe difficulties to overcome in making in situ measurements, especially since a new variable, pressure, must be considered (Jannasch 1985). In order to measure the activity of autotrophic bacteria at these sites, 14CO2 fixation was measured in syringes incubated on the seabed (approximately 250 atm, 3  C) and on board ship (1 atm) a 3  C and 23  C. Little or no difference was found between the two samples incubated

Colorless Sulfur Bacteria

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at 3  C, and the bacteria responsible for the 14CO2 fixation were thus obviously barotolerant rather than barophilic. Moreover, 14 CO2 fixation sharply increased if thiosulfate was added, or when the samples were incubated at 23  C, indicating that mesophilic colorless sulfur bacteria were responsible (Tuttle et al. 1983; Wirsen et al. 1986).

coworkers (Høgslund et al. 2009) used a combination of microelectrodes, quantitative microautoradiography, and 15N-stable isotopes to infer the in situ physiology of Thioploca. From their results they concluded that Thioploca is well adapted to a fluctuating environment by accumulation and transportation of nitrate and sulfur.

Inferring the Activity of Colorless Sulfur Bacteria in Their Natural Habitat with Microelectrodes

Inferring the Activity of Colorless Sulfur Bacteria in Their Natural Habitat with Molecular Methods

A technique that has been used with some success in the study of in situ bacterial biofilms and immobilization for biotechnology employs the use of microelectrodes with a tip of 1 mm of less that can be progressively moved through a biolayer, gradually registering the gradients present. The slope of the gradient, combined with data on the diffusion coefficient for the substrate measured, can provide direct information on the flux and turnover of substrates and thus can give accurate information on in situ activities. The microelectrodes are frequently linked to a computer that not only controls the rate of passage of the electrode tip through the sediment or biofilm but also records and calculates the results (e.g., Revsbech et al. 1986). Among others, oxygen, pH, sulfide, carbon dioxide, and N2O microelectrodes have been used, but the use of some (e.g., sulfide, CO2) is limited by their low sensitivity at commonly used pH values. However, the oxygen electrode has been extensively used, especially in systems where photosynthesis is involved and oxygen supply can easily be controlled by modifying the availability of light (e.g., Jensen and Revsbech 1989; Revsbech and Ward 1984). The construction of microelectrodes suitable for microbiology, and their use in various ecosystems, was extensively reviewed by Revsbech and Jørgensen (1986), but they are now commercially available (e.g., see www.unisense.com). Their use, in conjunction with some of the other methods mentioned above, provides a means of measuring actual activities in gradients, rather than potential activities in in vitro cultures. Microelectrodes, alone or in combination with other approaches, have been used in several studies to infer the in situ physiology of the colorless sulfur bacterium Beggiatoa. Schulz-Vogt and coworkers, for instance, used microelectrodes for oxygen, sulfide, nitrate, and pH to study the chemotactic response of freshwater Beggiatoa on different concentrations of nitrate (Kamp et al. (2006). Preisler et al. (2007) used similar microsensors to study the ecological niche of nitrate-storing Beggiatoa in coastal sediments and their contribution in the removal of sulfide. Hinck and coworkers (2007) used a combination of different microsensors, stable isotopes, and molecular techniques to study the dial cycling of Beggiatoa in hypersaline microbial mats. Similar microsensor studies were carried out for other colorless sulfur bacteria. Zopfi et al. (2001) used microelectrodes for oxygen and nitrate to study nitrate and sulfur storage in Thioploca species collected from the Bay of Concepcio´n, Chile. They found positive chemotaxis of Thioploca after addition of nitrate and nitrite. In a subsequent study, Jørgensen and

Currently, different molecular approaches are being used to infer the in situ activity of microorganisms in general and colorless sulfur bacteria in particular. One of the approaches is MARFISH, which is a combination of microautoradiography (MAR) and fluorescence in situ hybridization (FISH). In this approach, a mixed microbial community is briefly incubated with a radioactive labeled substrate such as [14C]-acetate. Subsequently, samples are taken at different time intervals and fixed in paraformaldehyde. The specimens are incubated with fluorescently labeled probes and covered with a photographic emulsion. After development of the photograph, the specimens can be seen under the microscope as the bacteria that took up the labeled substrate are covered with silver grains. The identity of these bacteria can be determined by fluorescence in situ hybridization. Nielsen and coworkers used MAR-FISH to study the in situ physiology of Thiothrix in wastewater treatment plants. They could demonstrate that Thiothrix was very versatile, being able to consume acetate and/or bicarbonate under heterotrophic, mixotrophic, and chemolithoautotrophic conditions (Nielsen et al. 2000). In a later study, they developed quantitative microautoradiography (QMAR) and fluorescence in situ hybridization (FISH) to determine the quantitative uptake of specific substrates by Thiothrix in activated sludge and found that the substrate affinity (Ks) for acetate was 2.4 mM (Nielsen et al. 2003). To increase the fluorescence in in situ hybridization experiments, CARD-FISH (catalyzed reported deposition-fluorescence in situ hybridization) was used. In this approach, the oligonucleotide probe, which is labeled with the enzyme horseradish peroxidase, catalyzes the deposition of fluorescently labeled tyramide molecules at the site of the probe hybridization, resulting in a strongly enhanced fluorescent signal (Amann and Fuchs 2008). By using 16S rRNA gene cloning, qPCR, and a combination of catalyzed reporter deposition-fluorescence in situ hybridization (CARD-FISH) with microautoradiography after incubation with [14C]-bicarbonate, Grote and coworkers (2007, 2008) found that Epsilonproteobacteria, and in particular those closely related to Sulfurimonas, were the main organisms responsible for CO2-fixation in the sulfidic waters of the pelagic redoxclines of the Baltic and Black Seas. Another way to measure activity in situ is the use of substrates labeled with stable isotopes, such as [13C]-acetate. As a consequence, all molecules in the bacterium that consumed the substrate will be labeled with the ‘‘heavy’’ carbon. The nucleic acids are extracted, and the 13C-labeled DNA

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(‘‘heavy’’ fraction) can be separated from 12C-labeled DNA (‘‘light’’ fraction) by cesium chloride or cesium trifluoroacetate gradient centrifugation, because of their difference in mass. Subsequently, the separated DNAs can be further analyzed using PCR amplification combined with cloning, DGGE, or NGS. The approach is known as DNA-SIP (DNA-stable isotope probing) or RNA-SIP (see Neufeld et al. (2007) for an overview). By using rRNA-based stable isotope probing (RNA-SIP) on water samples incubated with [13C]-labeled bicarbonate, they could reveal that 2 Gammaproteobacteria and Sulfurimonas were responsible for feeding the microbial food web in the redoxcline of the central Baltic Sea (Glaubitz et al. 2009). By using a combination of 13C labeling, FISH and secondary ion mass spectrometry (SIMS), Dattagupta et al. (2009) could show a novel symbiosis between the chemolithoautotrophic Thiothrix sp. and the freshwater cave amphipod Niphargus ictus and demonstrated that Thiothrix sp. was growing autotrophically. Zhang and coworkers (2005) used confocal laser-scanning microscopy, lipid biomarkers, stable carbon isotopes, and 16S rRNA gene sequencing to infer carbon cycling within Beggiatoadominated microbial mats associated with gas hydrates and cold seeps in the Gulf of Mexico. Recently, Wendeberg and coworkers (2012) studied in situ gene expression of pmoA (encoding subunit A of the particular methane monooxygenase) and aprA (encoding the subunit A of the dissimilatory adenosine-50 -phosphosulfate reductase) in methane- and sulfur-oxidizing symbionts of the hydrothermal vent mussel Bathymodiolus puteoserpentis. They found the highest mRNA expression levels at the ciliated epithelium of the gills, indicating a rapid response of the cells to incoming seawater rich in methane, reduced sulfur compounds and oxygen.

Conclusion New insights into the pathways of sulfur metabolism in the colorless sulfur bacteria have done away with the old unifying concept of sulfur metabolism, as it is now clear that there are diverse pathways in the organisms investigated thus far. Due to the revolution in DNA sequencing, the genomes of many different colorless sulfur bacteria (e.g., Acidithiobacillus, Sulfolobus, Sulfurihydrogenibium, Sulfurimonas, Sulfuricurvum, Thioalkalimicrobium, Thioalkalivibrio, Thiobacillus, Thiomicrospira) are currently available or underway (see the Genomes OnLine Database (www.genomesonline.org; Pagani et al. 2012). With these sequences in hand it will now be possible to study the evolution and ecophysiology, including the pathways in sulfur metabolism, of colorless sulfur bacteria in great detail. Whitaker and coworkers, for instance, studied the biogeographical structure of the pan-genome of Sulfolobus islandicus by comparative analysis of seven S. islandicus genomes from three different locations (Reno et al. 2009). They found that there was no gene flow between the geographically isolated populations. Beller and coworkers (2006) used whole genome transcriptomics to study thiosulfate oxidation by Thiobacillus

denitrificans under aerobic versus denitrifying conditions. Genes that were upregulated under aerobic conditions were siderophore-related genes, cytochrome cbb3 oxidase genes, genes (cbbl, cbbS) encoding form I RuBisCO, and chaperone genes, while genes upregulated under denitrifying conditions included nar, nir, and nor, genes (cbbM) encoding form II RuBisCO, and genes involved in the oxidation of sulfur compounds (i.e., sqr and dsrC). Yamamoto et al. (2010) used transcriptomics to study the sulfur metabolism in Sulfurovum sp. NBC37-1. In addition to whole genome sequencing of isolates, the use of the so-called meta-omics approach (i.e., metagenomics (e.g., Tringe et al. 2005), metatranscriptomics (e.g., Stewart et al., 2012), and metaproteomics (e.g., Siggins et al. 2012) makes it possible to study colorless sulfur bacteria that resist being isolated in pure culture. Although the ‘‘meta-omics’’ approach is very powerful, it is often difficult to obtain complete genomes from microbial communities that contain many different microorganisms. To circumvent this limitation, single cells can be isolated from environmental samples by micromanipulation or other means and their genomes can subsequently be sequenced (e.g., Yilmaz and Singh 2011; Martinez-Garcia et al. 2012). Mussman and coworkers (2007) used this approach to sequence the genome of single filaments of Beggiatoa. From these genomes, they could reconstruct pathways for sulfur oxidation, nitrate and oxygen respiration, and CO2 fixation confirming the chemolithoautotrophic physiology of Beggiatoa. Recently, Salman et al. (2011) handpicked individual cells of the large colorless sulfur bacteria Thiomargarita namibiensis, Thioploca araucae, and Thioploca chileae and sequenced their 16S rRNA genes and internal transcribed spacer (ITS) regions to determine their classification. However, a logical next step would be to sequence their complete genomes. In addition, high-throughput cultivation techniques and methods mimicking environmental parameters more accurately, such as gradient systems and in situ cultivation, are currently used to increase the success rate of isolation of bacteria in general and colorless sulfur bacteria in particular (see Alain and Querellou (2009) for an overview). Isolation of strains in pure culture is still essential to obtain a comprehensive understanding of the (eco)physiology of the bacteria. Since the writing of the previous edition of this chapter, one important question has been answered – should the colorless sulfur bacteria still be considered a taxonomic group? As discussed throughout this paper, the use as a taxonomic criterion of the ability to gain energy from the oxidation of inorganic reduced sulfur compounds has resulted in the definition of a very heterogeneous group, collectively known as the colorless sulfur bacteria. The possession of the relevant pathways for growth on reduced sulfur compounds is of no greater taxonomic relevance than the ability to use the Calvin cycle or to grow on hydrogen. As the results obtained with RNA and DNA analysis, have confirmed, we are seeing the result of evolutionary convergence towards the (eco)physiological properties encountered in many of the colorless sulfur bacteria. The extreme heterogeneity of the group is further emphasized as other long-known bacteria are tested and found to also possess the properties of colorless

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sulfur bacteria. Indeed, the common lack of a test for thiosulfate or sulfide oxidation in routine taxonomic screening has meant that the sulfur-oxidizing potential of species of genera such as Paracoccus, Pseudomonas and Alcaligenes are only now being recognized. That said, despite their morphological and phylogenetic diversity, the colorless sulfur bacteria present a coherent picture in physiological terms. As it is generally the physiological specifications of an organism that define its ecological significance, the reclassification of the colorless sulfur bacteria may present something of a microbiological dilemma because new taxonomic relationships bear little relation to the ecophysiological activities of the organisms. Thus, in spite of the reallocation of species among different genera, research can only profit from the retaining of physiological as well as taxonomic groupings, such as the sulfate reducers, nitrogen fixers, denitrifiers, and colorless sulfur bacteria.

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16 Bacterial Stress Response Eliora Z. Ron Department of Molecular Microbiology and Biotechnology, The George S. Wise Faculty of Life Sciences, Tel Aviv University, Tel Aviv, Israel

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 589 Global Regulatory Networks in Bacteria . . . . . . . . . . . . . . . . . . 589 Stress Response, Stimulons, and Regulons . . . . . . . . . . . . . . . . . 590 The Heat-Shock Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 590 Heat-Shock Control Elements in Gram-Negative Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 592 Sigma-32-Controlled Genes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 592 Genes Controlled by sE . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 593 HrcA-CIRCE-Controlled Genes . . . . . . . . . . . . . . . . . . . . . . . . . 594 Induction of Heat-Shock Genes by Changes in Ribosome Accessibility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 595 ROSE-Controlled Genes and Thermosensors . . . . . . . 595 E. coli HNS-Controlled Genes . . . . . . . . . . . . . . . . . . . . . . . 595 Additional Regulatory Elements . . . . . . . . . . . . . . . . . . . . . . . . . 595 Regulation by Modulating Transcript Stability . . . . . . . . . 595 The General Stress Response in E. coli . . . . . . . . . . . . . . . . . . . . 595 Control of the Heat-Shock Response and the General Stress Response in Gram-Positive Bacteria . . . . . . . . . . . . . . . . 596 Heat-Shock Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 597 The General Stress Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . 597 Complexity of the Stress Response Networks . . . . . . . . . . . . . . 598 Abstract Bacteria respond to stress by regulatory networks which modulate gene expression. These response mechanisms are essential for coping with the stress and for adapting to the new conditions. The regulation of the stress response involves several molecular pathways which control transcription, translation, and stability of transcripts and of proteins. These molecular responses are the topic of this chapter, which focuses on adaptation to upshift in temperature (heat-shock response) and to starvation-stationary conditions (general stress response).

Introduction Most bacteria live in a dynamic environment where temperature, availability of nutrients, and presence of various chemicals vary. Quick adaptation to these environmental changes is

carried out by a series of global regulatory networks that control the simultaneous expression of a large number of genes. There are global regulatory systems that respond to change of temperature, pH, nutrients, salts, and oxidation. The level of response by these regulatory networks is proportional to the extent of the change. Since the response level is highest under changes that constitute a stress condition, the control networks are labeled ‘‘stress response’’ systems. The stress response systems show a high degree of similarity in prokaryotes, and some (e.g., the heat-shock response) are also conserved in eukaryotes and archaea. However, the conditions under which the response systems are activated differ significantly from one organism to another. Clearly, the temperatures in which the heat-shock response is activated will be much lower for a mesophile than for a thermophile, or the response to salt stress will be completely different in halophiles.

Global Regulatory Networks in Bacteria The first attempts to study the extent of such regulatory networks were based on proteomic analysis, using O’Farrell two-dimensional (2D) gels, and resulted in the identification of the large group of Escherichia coli heat-shock proteins (Neidhardt et al. 1981; O’Farrell 1975). Later, proteomic-based experiments followed by microarray studies of gene transcription (Hatfield et al. 2003) revealed the size and composition of the various stress-induced stimulons of E. coli (RA et al. 1987). This induction of large groups of genes in response to a specific environment suggested the existence of global regulatory systems that control the expression of large regulons. Gene expression can be regulated at the level of transcription or posttranscription. The level of transcription can be regulated by positive control elements—activators—or by negative control elements—repressors. Some of these control elements are specific for one gene, whereas others control a large group of genes, thus creating a regulon. In addition to transcriptional regulation, many posttranscriptional regulatory systems evolved, affecting different steps along the way from the gene to the active protein. The posttranscriptional regulatory systems control the stability of the mRNA and the rate of translation initiation. In addition, they can determine the stability of the protein and its activity by carrying out posttranslational modifications. The existence of all of the control elements described here was demonstrated in the global regulatory systems that control the response to heat shock and other environmental and physiological conditions.

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_79, # Springer-Verlag Berlin Heidelberg 2013

590

16

Bacterial Stress Response

Transcriptional regulation is the primary mechanism that regulates gene expression. The process of RNA synthesis and its control were extensively studied in bacteria, especially in E. coli and Bacillus subtilis (Burgess and Anthony 2001). The E. coli DNA-dependent RNA polymerase is the enzyme responsible for all cellular RNA synthesis. This enzyme consists of a core (subunits a2bb0 o) that is capable of elongation and termination of transcription and an additional subunit (s) which binds to the RNA polymerase to form the holoenzyme, increases the efficiency of transcription initiation, and determines specific promoter recognition (Burgess et al. 1969). In E. coli, there are seven known sigma factors: s70 (sigma 70) the vegetative sigma factor and sS, s32, sF, sE, sfecI, and s54 (Burgess and Anthony 2001; Helmann and Chamberlin 1988; Lonetto et al. 1992). The sigma factors serve as master regulators mainly by competition for the core RNA polymerase, which is the limiting component of the transcription machinery (Ishihama 2000). Additional regulation of transcription is exerted by repressors, transcriptional activators, sigma-binding anti-sigma factors, and even by small RNAs (Helmann 1999; Hughes and Mathee 1998; Ishihama 2000; Narberhaus et al. 2009; Severinov 2000; Storz et al. 2011; Vicente et al. 1999; Vogel and Luisi 2011; Wassarman and Storz 2000). In the last years, it became clear that two additional control mechanisms are involved in bacterial response to environmental stress—proteolysis (Gur et al. 2011; Hengge 2009; Schmidt et al. 2009) and modulation of transcript stability (Caron et al. 2010; Henkin 2009; Rasouly et al. 2007; Shenhar et al. 2009). These various control elements regulate the expression of genes during environmental conditions such as starvation, sporulation, and additional stress conditions. For example, the E. coli stationary phase is regulated by the master regulator sS (Lange and Hengge-Aronis 1991). The levels of sS itself are affected by cis and trans elements—small molecules, such as guanosine 50 diphosphate 30 -diphosphate (ppGpp) and homoserine lactone, and the proteins that react to them, such as cAMP receptor protein (CRP)-cAMP (Hengge-Aronis 2000). Sigma factor S regulates the induction of more than 50 genes, and its expression involves small RNA and Hfq (Gottesman et al. 2006; Sittka et al. 2008; Vogel and Luisi 2011; Zhang et al. 2003) (Weber et al. 2005). All of these elements create a complex regulatory network that enables the bacterial cell to adapt to the changing environment.

Stress Response, Stimulons, and Regulons In bacteria, the stress responses are regulated by several control patterns: (1) Transcriptional control by alternative sigma factors is the most prevalent control pattern. Basically, genes or operons that belong to a specific response regulon contain a promoter that is recognized by a specific, alternative, sigma factor. The function of this sigma factor correlates with the conditions that bring about the response. As an example, in Gram-negative bacteria, the response to elevated temperatures is mediated by two alternative sigma factors (s32 and sE) whose activities are temperature dependent. (2) Transcription is controlled by

repressor binding to a DNA control element. An example of this control is the HrcA repressor that binds to a conserved inverted repeat control element known as ‘‘CIRCE’’ (for ‘‘controlling inverted repeat of chaperone expression’’) present upstream of operons that code for heat-shock proteins in Gram-positive and some Gram-negative bacteria. (3) Transcription is controlled by proteolysis. Well defined is the SOS (This is the real SOS—save our souls—and not ‘‘salt overly sensitive’’) response to genotoxic effects, which is mediated by a series of autoregulated proteases. Recently, control by proteolysis has emerged as one of the major systems regulating the availability of alternative sigma factors and other stress-related global processes (Jenal and Hengge-Aronis 2003). (4) Transcription is controlled by small RNAs. Recent findings indicate that small RNAs, about 50 of which are present in the Escherichia coli genome, control the cellular concentration of RpoS (sigma38), the alternative sigma factor of the starvation (or stationary) response. Small RNAs also control the response to oxidative stress and are involved in pathogenesis (Basineni et al. 2009; Frohlich and Vogel 2009; Papenfort et al. 2009; Podkaminski and Vogel 2010). A regulon is defined as all genes regulated by the same control pattern, while a stimulon is defined as all the genes whose expression responds to the same conditions. Stimulons are easily delineated by monitoring gene expression in a microarray or on two-dimensional protein gels. Regulons can only be established following characterization of the molecular basis for the change in gene expression. Clearly, level of overlap between the various regulons and stimulons is high. Thus, the stimulon that responds to shifts to higher temperatures contains genes from at least two regulons (i.e., s32 and sE). Yet, some of the genes of the s32 regulon may also be controlled by the HrcA repressor and so on. Here, the focus is on two stress response networks—one responding to shifts to higher temperatures (heat-shock response) and the other to limitation of carbon source and stationary phase (general stress response).

The Heat-Shock Response The heat-shock response was the first global regulatory system to be discovered and is one of the most fundamental. This response is general, found in all living cells examined (Craig 1985), and is a protective and homeostatic cellular process that increases thermotolerance. It has been studied in many cellular systems such as bacteria, yeast, insects (Drosophila melanogaster; (Michaud et al. 1997)), worms (Caenorhabditis elegans (Rose and Rankin 2001)), and mammals (Srivastava 2002). The heatshock response is characterized by the induction of a large set of proteins (heat-shock proteins—HSPs) as a result of a rapid increase in the environmental temperature. Many of the HSPs are molecular chaperones (e.g., GroEL, GroES, DnaK, and DnaJ) and ATP-dependent proteases (e.g., ClpP, Lon (La), and HslVU) that play a critical role in the restoration of protein folding and in protein degradation under normal and stress conditions.

Bacterial Stress Response

Proteins such as GroEL (the bacterial homolog of Hsp 60) and DnaK (the bacterial homolog of Hsp 70) are highly conserved in evolution all the way from bacteria to humans (Boorstein et al. 1994; Gupta 1995). Although the major proteins in the heatshock response are highly conserved, the regulation of the response varies between different organisms and different bacterial species. Several regulatory systems evolved in bacteria and will be discussed here. The Hsps are important for protection against environmental stress, and they produce tolerance against high temperature, high salt, and heavy metals (Hecker and Volker 1998; Inbar and Ron 1993; Val and Cronan 1998; RA et al. 1987). Heat-shock proteins also play critical roles in bacterial virulence and in protective systems such as the human immune system (Hecker and Volker 1998; Inbar and Ron 1993; Val and Cronan 1998; RA et al. 1987). Several Hsps were found to protect against damage induced by temperature upshifts. Among the characterized proteins are the main cellular chaperone machineries GroE and DnaK, the ATP-dependent proteases Lon (La), HslVU, ClpP, DegP, and FtsH (HflB), and other proteins involved in protein

16

folding, refolding, quality control, and degradation. GroE and DnaK are both multimeric complexes that have ATP-dependent activity (Kandror et al. 1994; Sherman and Goldberg 1992, 1996). The GroE catalytic complex involves GroEL and GroES in a ratio of 1:2, creating a football-shape molecular structure (Sparrer et al. 1997). This complex catalyzes protein refolding and is involved in protein degradation by the ATP-dependent proteases (Kandror et al. 1994; Sherman and Goldberg 1992, 1996). These ATP-dependent proteases degrade abnormal proteins under stress and nonstress conditions and in addition play major regulatory functions by controlling the degradation of specific proteins (Deuerling et al. 1997; Goldberg 1972; Gottesman 1996; Maurizi 1992; Zhou et al. 2001). The role of these and other E. coli Hsps in protection against temperatureinduced damage is summarized in > Table 16.1. Heat shock—a rapid upshift in the environmental temperature—results in various physical and chemical changes in bacterial proteins and membranes. Presumably, these changes, such as protein unfolding, are detected by cellular systems, which induce the large set of heat-shock proteins to cope with the

. Table 16.1 Major heat-shock proteins of Escherichia coli Protein

Function

Molecular weight (kDa)

Theoretical pI

References

ClpB

Chaperone

96

5.37

Kitagawa et al. (1991)

DnaJ

Chaperone

39

7.98

Bardwell et al. (1986)

DnaK

Chaperone

69

4.83

Bardwell and Craig (1984)

GroEL

Chaperone

57

4.85

Neidhardt et al. (1981)

GroES

Chaperone

10

5.15

Tilly et al. (1983)

HslR (Hsp15)

Chaperone

15

9.94

Chuang and Blattner (1993)

Hsp33 (HslO)

Chaperone

33

4.65

Chuang and Blattner (1993)

HtpG

Chaperone

71

5.09

Bardwell and Craig (1987)

IbpA (HtpN, HslT)

Chaperone

16

5.57

Allen et al. (1992)

IbpB (HtpE, HslS)

Chaperone

16

5.19

Allen et al. (1992)

ClpP

Protease

24

5.52

Maurizi et al. (1990a)

ClpX

Protease

46

5.24

Gottesman et al. (1993)

DegP (HtrA)

Protease

50

8.65

Lipinska et al. (1988)

FtsH (HflB)

Protease

71

8.91

Herman et al. (1995a)

HslU (ClpY, HtpI)

Protease

49

5.24

Chuang and Blattner (1993)

HslV (ClpQ, HtpO)

Protease

19

5.96

Chuang and Blattner (1993)

Lon (La)

Protease

87

6.01

Gayda et al. (1985)

s32 (RpoH, HtpR, Hin, Fam)

Sigma factor

32

5.64

Landick et al. (1984)

s70 (RpoD, Alt)

Sigma factor

70

4.69

Burton et al. (1981)

24

s (s , RpoE)

Sigma factor

22

5.38

Raina et al. (1995)

PrpA (PphA)

Phosphatase

25

6.94

Missiakas et al. (1993)

HtpX

Unknown

32

6.60

Kornitzer et al. (1991)

E

HtpY (HtgA)

Unknown

21

9.44

Missiakas et al. (1993)

HtrC

Unknown

21

9.33

Raina and Georgopoulos (1990)

PspA

Unknown

25

5.39

Jovanovic et al. (1996)

FtsJ

Unknown

23

9.44

Herman et al. (1995a)

591

592

16

Bacterial Stress Response

changes and the potential damage. This heat-shock response is regulated by several control elements, thus dividing the major stimulon of heat-shock proteins into several regulatory groups (regulons). The heat-shock proteins are highly conserved, whereas the control of their expression is highly variable between organisms and even between various bacteria. One of the control elements found in Gram-negative bacteria is a heat-shock s factor that regulates the transcription of the major Hsps. The Gramnegative E. coli is a good example of this system because the synthesis of the major Hsps is regulated by the alternative sigma factor called ‘‘s32.’’ In addition, there is a group of proteins induced under conditions of elevated temperature that is regulated by another heat-shock sigma factor, sE (encoded by rpoE). In other Gram-negative bacteria, such as the Agrobacterium tumefaciens of the Alphaproteobacteria, the control systems are more complicated. For example, the transcription of GroESL synthesis is stimulated during heat shock by a s32-like activator, but in non-heat-shock conditions, transcription is repressed by the HrcA protein that binds to the CIRCE sequence upstream of the promoter region (Nakahigashi et al. 1999; Segal and Ron 1993). The control system of HrcA-CIRCE was first described in the Gram-positive Bacillus subtilis (Zuber and Schumann 1994). The following sections will describe the specific control mechanisms in various bacterial groups. In short, the heat-shock response in bacteria is controlled by one or a combination of both of the following control systems: (1) The first system involves alternative sigma factors that act as transcriptional activators by recognizing specific heat-shock promoters upstream of heat-shock genes. Among these are s32 and sE of the Gramnegative bacteria and sB of the Gram-positive bacteria. (2) The second system utilizes transcriptional repressors. The most conserved and the most ubiquitous among these repressors is HrcA (heat regulation at CIRCE), which binds to a conserved CIRCE present upstream of the heat-shock operons. Heat-shock operons controlled by HrcA-CIRCE are transcribed by the vegetative sigma factor sA (=s70) in Gram-positive bacteria and by the heat-shock sigma factor s32 in Gram-negative bacteria.

Heat-Shock Control Elements in Gram-Negative Bacteria The first model organism for studying the heat-shock response in Gram-negative bacteria was E. coli. Most of the heat-shock genes of this bacterium are regulated by transcriptional activators, the alternative sigma factors (s32 or sE).

Sigma-32-Controlled Genes The heat-shock response of Gram-negative bacteria is regulated mainly by the alternative sigma factors s32 and sE (Morita et al. 2000). Sigma 32 is a master regulator encoded by the rpoH (htpR or hin) gene that was the first of a group of minor sigma factors

discovered in E. coli (Grossman et al. 1984; Landick et al. 1984; Yuzawa et al. 1993). This discovery of minor sigma factors led to the general concept of gene regulation by specific sigma-factordependent transcription. In E. coli, at least 34 heat-shock genes (out of 51 heat-shock-induced loci) are regulated by s32 (Morita et al. 2000; Richmond et al. 1999). The genes were identified by transcription analysis of specific genes, an examination of the synthesis rates of individual proteins, or proteomics and genomics approaches. This regulon includes all the major cytoplasmic Hsps of E. coli. The s32 regulon includes most of the proteins involved in protein folding, repair, and degradation. Such proteins are the heat-shock-induced molecular chaperones ClpB, DnaK, DnaJ, GroEL, and GroES, which are involved in protein folding and prevention of protein aggregation (Bardwell and Craig 1984; Bardwell et al. 1986; Kitagawa et al. 1991; Neidhardt et al. 1981; Tilly et al. 1983; Tomoyasu et al. 2001). The regulon comprises also all of the important cytosolic proteases Lon (La), ClpP, ClpX, HslV (ClpY), and HslU (ClpQ); (Chuang and Blattner 1993; Gayda et al. 1985; Goldberg 1972; Maurizi et al. 1990b) and the membranal metaloprotease FtsH (HflB) (Herman et al. 1995a, b; Tomoyasu et al. 1995). Other important s32-regulated proteins are HTS (homoserine transsuccinylase), which is a key enzyme in methionine biosynthesis (Biran et al. 1995), proteins involved in protein isomerization (PpiD (Dartigalongue and Raina 1998) and HtrM (Raina and Georgopoulos 1991)), and the vegetative sigma factor (s70) (Burton et al. 1981). Recently, a new group of heat proteins was identified. These are conserved heat-shock proteins essential for growth at high temperature and involved in regulating and maintaining efficient translation under heat shock (Rasouly et al. 2007; Rene and Alix 2011). This group includes (FtsJ) (Bugl et al. 2000; Caldas et al. 2000a, b), fidelity of translation (YciH and FtsJ) (Lomakin et al. 2006; Widerak et al. 2005) (Bucca et al. 1995; De Las Penas et al. 1997), and the recycling of 50S particles following aberrant translation termination events (Hsp15) (Korber et al. 2000, 1999). Homologs of rpoH were identified in more than 20 species of Eubacteria from the alpha, beta, and gamma subgroups of Proteobacteria (Andersson et al. 1998); (Huang et al. 1998); (Karls et al. 1998); (Emetz and Klug 1998; Nakahigashi et al. 1998, 1999, 2001; Narberhaus et al. 1997; Sahu et al. 1997). In some of these bacteria, the rpoH homologs demonstrate translational induction and stabilization upon heat shock, which are very similar to those found in E. coli (Nakahigashi et al. 1998). The general function of the s32 regulon was studied in several bacterial species by analysis of rpoH mutants. These mutants were usually found to be temperature sensitive (Huang et al. 1998; Nakahigashi et al. 1999; Zhou et al. 1988). As expected from their temperature-sensitive phenotype, some of the heat-shock proteins are essential at elevated temperature. The levels of s32 and its activity are temperature regulated at several levels. At low temperature (30  C), when low amounts of heat-shock proteins are required, the intracellular concentration

Bacterial Stress Response

of s32 is fewer than 50 molecules per cell (Craig and Gross 1991; Straus et al. 1987). These low levels are maintained due to transcriptional repression and protein instability. Upon a rapid shift to 42  C, the level increases 15–20-fold within 5 min and then declines to a new steady state level, two–threefold higher than the pre-shift level (Straus et al. 1987). The levels and the time course of s32 induction are sufficient for the necessary induction of heat-shock-gene expression upon temperature upshift. A relatively modest heat shock activates the translation of rpoH transcripts and transiently stabilizes s32 (Nagai et al. 1991a, b; Straus et al. 1987), whereas a severe heat shock (a rapid shift from 30  C to 50  C) can also activate rpoH transcription (Morita et al. 2000). The decrease in the synthesis of heat-shock proteins upon temperature downshift is primarily a result of the decrease in s32 activity (rather than its levels) caused mainly by an excess of the DnaK chaperone machinery (Straus et al. 1989; Taura et al. 1989). The transcriptional regulation of the rpoH gene is very complex. It can be transcribed from at least four promoters, three of them (P1, P4, and P5) are recognized by the vegetative s70, and the fourth (P3) is recognized by sE (Erickson et al. 1987; Nagai et al. 1990). P3 and P4 transcription of rpoH is negatively regulated by DnaA (Wang and Kaguni 1989), and P4 and P5 transcription is controlled by an additional negative control system—the cAMP-CRP/CytR nucleoprotein complex (Kallipolitis and Valentin-Hansen 1998). Several findings indicate that the heat-shock-induced s32 levels are also regulated at the translational level. Expression of rpoH-lacZ translational fusion but not transcriptional fusion can be induced. Furthermore, heat induction of the fusion protein occurs even when RNA synthesis is inhibited (Nagai et al. 1991a, b). Recent results based on extensive in vivo and in vitro experiments related to the secondary RNA structure have shown that the translation regulation of RpoH is mediated by the rpoH mRNA’s secondary structure (Morita et al. 1999a, b, 2000). Sigma 32 level is regulated by not only its expression level but also the turnover of the protein. Although this protein is unstable during normal growth at 30  C (or even at 42  C), significant stabilization occurs immediately upon temperature upshift from 30  C to 42  C and continues for 4–5 min (Straus et al. 1987). The protein instability involves the DnaK chaperone machinery. Mutants in DnaK, DnaJ, or GrpE markedly stabilize s32 under nonstress conditions (Straus et al. 1990; Tilly et al. 1983, 1989), indicating this involvement of these proteins in s32 turnover. The initial studies suggested that the membrane-associated metalloprotease FtsH (HflB) is responsible for s32 degradation (Herman et al. 1995a; Tomoyasu et al. 1995). However, later studies were able to show that the cytosolic proteases Lon (La), HslVU, and ClpP are also involved in s32 degradation (Kanemori et al. 1997, 1999; Morita et al. 2000; Wawrzynow et al. 1995). Although the relative significance of each protease is difficult to determine in s32 degradation, the latter three proteases seem to play a significant role in the degradation, possibly even equivalent to that of FtsH (Kanemori et al. 1997). Presumably during heat shock, the DnaK machinery and the proteases

16

become occupied by the misfolded and unfolded proteins that accumulate because of the denaturing effect of temperature increase. Consequently, levels of the proteolytic machinery are insufficient to bring about s32 degradation, and it accumulates and activates the transcription of the heat-shock genes. Since the DnaK chaperones and the proteases have s32 promoters, their synthesis is increased, and therefore a few minutes after the temperature upshift, the level of the proteases and chaperones is high enough to destabilize s32, bringing the level of the heatshock proteins to a new steady state. The final level of s32 regulation is activity regulation (Morita et al. 2000). This regulation operates mainly by creating ternary complexes of (DnaK-ADP)-DnaJ-s32 that sequester the s32 that competes with the RNA polymerase core enzyme (Gamer et al. 1992, 1996; Liberek et al. 1992; Liberek and Georgopoulos 1993). Then, GrpE binds to the ternary complex and stimulates ADP release to allow rebinding of ATP. This cycle of binding and release appears to play an important role in s32 activity (and possibly stability) in vivo (Gamer et al. 1992; Morita et al. 2000). The s32 control system has been well characterized in E. coli and other Gammaproteobacteria. However, s32-like heat-shock transcriptional activators have recently been demonstrated in other bacteria, such as Agrobacterium tumefaciens of the Alphaproteobacteria (Nakahigashi et al. 1995, 1998a, 1999; Segal and Ron 1995a, 1996b). The s32 of the Alphaproteobacteria is different from that of E. coli, and the heat-shock promoters are also different in the two groups of Gram-negative bacteria (Nakahigashi et al. 1999; Segal and Ron 1995a, 1996b). The physiological difference of the two sigma factors may be more important: while the E. coli s32 is unstable and tightly controlled by proteolysis carried out by the FtsH protease, the alphaproteobacterial s32 is a stable protein, whose activity is affected mainly by a DnaK-mediated control (Nakahigashi et al. 2001) (> Fig. 16.1).

Genes Controlled by sE Another alternative sigma factor involved in the heat-shock response is sE (s24), which was found to be an essential gene in E. coli at all temperatures (De Las Penas et al. 1997). Presumably, the sE regulon protects cells against extracytoplasmic stress-derived damage. Genes belonging to the sE regulon are important for bacterial pathogenesis: the mucoid phenotype of Pseudomonas aeruginosa in cystic fibrosis infections is controlled by AlgU, an analogue of sE (Yu et al. 1995), and rpoE mutants of Salmonella typhimurium are defective in growth inside cells (Humphreys et al. 1999). The E. coli sE controls the expression of genes encoding periplasmic folding catalysts, proteases, biosynthetic enzymes for the lipopolysaccharide component lipid A, and other proteins whose functions are involved with the cell envelope. Members of this regulon include periplasmic proteins that are involved in protein metabolism: the protease DegP (HtrA)

593

594

16

Bacterial Stress Response

Putative promoters Heat shock promoter in a-subdivision

CTTG

CYTAT-T

Heat shock promoter in g-subdivision

TCTC-CCTTGAA

CCCAT-AT

TTGACA

TATAAT

Vegetative promoter in g- and a-subdivisions

Promoter recognition domains 4.2

2.4

Sigma factor domain

a-Subdivision g -Subdivision

s-32 s-32

IKASIQEYILRSWSLVKMGTT IKAEIHEYVLRNWRIVKVATT ∗ ∗ ∗ ∗ ∗∗

a-Subdivision g -Subdivision

s-70 s-70

IRQAITRSIADQARTIRIPVHM IRQAITRSIADQARTIRIPVHM

YGVSRERVRQIEKRAMKKLR YGVSAERVRQLEKNAMKKLR ∗ ∗ ∗ FSVTRERIRQIEAKALRNVK FDVTRERIRQIEAKALRNVR ∗ ∗

. Fig. 16.1 Putative heat-shock promoters and promoter recognition domains of s-32 and s-70 in alpha-purple and gamma-purple proteobacteria (Nakahigashi et al. 1999; Segal and Ron 1995a)

and the periplasmic peptidyl-prolyl isomerase FkpA (Dartigalongue and Raina 1998; Erickson and Gross 1989; Strauch et al. 1989). As mentioned above, sE activates transcription of rpoH under conditions of severe heat shock, and because it has a sE promoter, it also regulates itself. The response is regulated by RseA (an inner membrane antagonist of sE), RseB (a periplasmic protein that binds to the periplasmic face of RseA), and the proteases DegS and YaeL. Envelope stress promotes RseA degradation, which occurs by a proteolytic cascade initiated by DegS. There is evidence that one sE-inducing stress (OmpC overexpression) directly activates DegS to cleave RseA (Alba and Gross 2004).

HrcA-CIRCE-Controlled Genes The HrcA-CIRCE repression system is the major system regulating the operons coding for chaperones in Gram-positive bacteria. This system is comprised of an inverted repeat cis element and a trans protein repressor encoded by the hrcA gene. The first reported inverted repeat upstream to the groE operon was found in Mycobacterium tuberculosis in 1989 (Baird et al. 1989). Recognition of this element as a widespread heat-shock control element for the groE and dnaK operons took several years. Several lines of direct evidence for the role of CIRCE as a negative cis element were obtained (Narberhaus 1999): (1) deletion of the inverted repeat relieved the repression of a reporter gene fusion (amyS); (van Asseldonk et al. 1993), (2) placement of CIRCE behind a foreign promoter reduced the expression of the downstream gene (Zuber and Schumann 1994), and (3) site-directed mutation, or the removal of three or four nucleotides in one arm of the inverted repeat, resulted in an elevated transcription of the downstream genes at normal

growth temperature (Babst et al. 1996; Zuber and Schumann 1994). Transcription remained derepressed when the inverted repeat was restored by compensating mutations in the second arm of the inverted repeat. Therefore, the CIRCE is not only a potential stem and loop structure (because its sequence by itself is required for repression) but also a binding site for a sequence-specific repressor protein that binds to the CIRCE. The elucidation of CIRCE as a potential repressor-binding site initiated a search for the counterpart repressor. Major steps toward tracking the repressor were accomplished by two observations (Narberhaus 1999): (1) a deletion of orf39—the first gene of the dnaK operon of B. subtilis resulted in an elevated levels of groE transcript (Schulz et al. 1995), and (2) B. subtilis mutants affected in the regulation of groE and dnaK operons were mapped to orf39 (Yuan and Wong 1995b). Moreover, production of Orf39 from a plasmid that carries a functional copy of orf39 restored the repression activity in one of the mutants (Yuan and Wong 1995a). The binding of Orf39 to CIRCE was shown by gel retardation (Narberhaus 1999), and the name ‘‘HrcA’’ (‘‘heat regulation at CIRCE’’) was given to this protein after disruption of the equivalent gene in Caulobacter crescentus (Roberts et al. 1996). For several years, the HrcA-CIRCE system was found only in Gram-positive bacteria and was considered as a Gram-positive heat-shock control element. However, since the first discovery of the CIRCE element in the Gram-negative Alphaproteobacterium A. tumefaciens (Segal and Ron 1993), many CIRCE elements were identified in other Gram-negative bacteria. The inverted repeat was detected in a large number of phylogenetically distant bacteria, including Gram-negative bacteria of the Alpha, Beta, and Gamma1 purple Proteobacteria. The only groups where it is probably not present at all are the Gamma2 and Gamma3 purple bacteria, which also include the Gramnegative model organism E. coli (Ron et al. 1999; Segal and

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Ron 1998). The inverted repeat (TTAGCACTC-N9GAGTGCTAA) is highly conserved in all of the studied genes (Segal and Ron 1996a, 1998). In contrast to Gram-positive bacteria where CIRCEregulated genes are transcribed with the vegetative sigma factor (sA), in A. tumefaciens the groEL operon is HrcA-CIRCE controlled, but is transcribed mainly by s32 (Nakahigashi et al. 1999). In A. tumefaciens, it was possible to show, using 2D gels, that GroE proteins are the only proteins whose synthesis is repressed by the HrcA-CIRCE system (Rosen et al. 2002). In Bradyrhizobium japonicum, two groESL operons were found: groESL1 is s32 regulated, while groESL2 is CIRCE-HrcA-s96 dependent (s96 recognizes the housekeeping promoter of B. japonicum) (Munchbach et al. 1999a, b). The control of chaperone expression by the HrcA-CIRCE system seems to be more ancient than the s32-dependent transcription of heatshock genes because it is found in all the bacteria except two small groups that lost it during evolution, whereas s32-dependent transcription is found only in Gram-negative bacteria (Ron et al. 1999).

Additional Regulatory Elements

Induction of Heat-Shock Genes by Changes in Ribosome Accessibility

Regulation by Modulating Transcript Stability

ROSE-Controlled Genes and Thermosensors Expression of at least ten genes in B. japonicum, seven of which code for small Hsps, is under the control of ROSE (repression of heat-shock gene expression); (Munchbach et al. 1999a, b; Narberhaus et al. 1998). This negatively cis-acting DNA element confers temperature control to a s70-type promoter. ROSE elements are not restricted to B. japonicum but are also present in Bradyrhizobium sp. (Parasponia), Rhizobium sp. strain NGR234, and Mesorhizobium loti (Nocker et al. 2001a, b). The latest model for ROSE activity suggests that ROSE controls heat-shock protein expression by a temperature-dependent secondary structure of ROSE mRNA that controls the access of the ribosome to the ribosome binding site (Nocker et al. 2001a, b). These studies are an example of regulation of translation efficiency by RNA ‘‘thermometers’’ (Klinkert and Narberhaus 2009; Waldminghaus et al. 2009).

E. coli HNS-Controlled Genes Heat-shock protein 31 (Hsp31), the product of the hchA (yedU) gene, is a conserved heat-shock chaperone (Mujacic et al. 2004; Sastry et al. 2002). Recent results indicated that the level of Hsp31 is induced upon temperature upshifts even in rpoH mutants lacking s32. It was shown that the hchA gene is transcriptionally regulated by the RpoS transcriptional activator, which regulates the general stress response in E. coli (Akbar and Price 1996; Andersson et al. 1998; Antelmann et al. 1997b), and that the heat-shock induction is presumably due to derepression by HNS (Andersson et al. 1998).

Proteome analysis of A. tumefaciens and in its mutants deleted for rpoH, hrcA, or in both showed that the heat-shock induction of 32 (out of 56) heat-shock proteins is independent of RpoH and HrcA. These results indicate the existence of additional regulatory factors in the A. tumefaciens heat-shock response (Rosen et al. 2001, 2002). These uncharacterized regulatory elements may also involve ROSE because A. tumefaciens belongs to the Rhizobiaceae group. An additional unique posttranscriptional control mechanism demonstrated in A. tumefaciens involved a specific cleavage of the groESL operon transcript. The resulting groES transcript is rapidly degraded, whereas the groEL transcript is stable, leading to a differential expression of the two genes of the operon—as could be detected by quantitative analysis of the protein expression, using 2D gels (Rosen et al. 2002; Segal and Ron 1995b). This mRNA processing is temperature dependent and constitutes the first example of a controlled processing of transcripts in bacteria.

A novel regulatory mechanism which is important in stress response is modulation of transcript stability. Thus, during heat shock, the transcript stability of the rpoH transcript is significantly increased (Shenhar et al. 2009), and a similar increase occurs for the rpoS transcript during stationary and starvation stress (Hengge-Aronis 2002). And, as already mentioned, stabilization of transcript also plays an important role in the regulation of expression of the YedU (HchA) gene and the small heat-shock proteins (Klinkert and Narberhaus 2009; Rasouly et al. 2007; Waldminghaus et al. 2009). Recent data demonstrate the existence of an additional regulatory system at the level of transcript stability—upon heat shock, the stability of transcripts of heat-shock genes is considerably reduced (Shenhar et al. 2009). This reduction is essential for the transient nature of the heat-shock response and contributes to balanced growth after an upshift in temperature.

The General Stress Response in E. coli The ‘‘general stress response’’ is induced during carbon starvation or entry into stationary phase. In E. coli, these conditions result in a variety of physiological and morphological changes that, presumably, ensure survival during periods of prolonged starvation. Although this general stress response was believed to involve the induction of 30–50 proteins (Lange and HenggeAronis 1991), this stimulon now appears to be much larger and involve almost 500 genes, most of which are induced by osmotic shock. About half are induced by stationary phase or acidic stress, and many are induced by more than one or all of these stresses (Hengge 2009). The general stress response is also important in quorum sensing (Schuster et al. 2004). The general

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stress response involves several regulatory elements—transcription of stress genes by an alternative sigma factor—RpoS— proteolysis and modulation of transcript stability. The genes coding for the general stress response in E. coli are transcribed by an alternative sigma factor, sS (RpoS), which recognizes a consensus promoter upstream of the general stress genes. The promoter specificity of sS has been difficult to determine, as the promoter it recognizes appears quite similar to those recognized by the vegetative s70. The specific sS promoter elements were recently characterized (Klauck et al. 2007; Singh et al. 2011; Typas et al. 2007a, b, c), and the results suggest that the selectivity is provided by the K173 (the lysine in position 173 of the amino acid sequence) in sS (which is glutamate in s70). sS binds to the C( 13) and the distal upstream (UP) element 35 of the promoter. The E. coli RpoS is a highly unstable protein, whose degradation is inhibited by various stress signals, such as carbon starvation, high osmolarity, and heat shock. As a consequence, these stresses result in the induction of sS-regulated stressprotective proteins (Bouche et al. 1998). Proteolysis of sS requires the response regulator RssB (a direct recognition factor with phosphorylation-dependent affinity for sS which targets sS to the ClpXP protease (Pruteanu and Hengge-Aronis 2002; Zhou et al. 2001)). Recognition of sS by the RssB/ClpXP system involves two distinct regions—region 2.5 of RpoS is a long a-helix which binds phosphorylated RssB. This binding exposes a second region of RpoS, located in the N-terminal part, which is a binding site for the hexameric ring of the ClpX chaperone (Studemann et al. 2003). Recent studies demonstrate the involvement of small, noncoding RNAs (Vogel et al. 2003) in the proteolysis of sS. These small noncoding RNA sequences are abundant—around 50 such sRNAs were described in E. coli. The levels of many of these sRNAs vary with changing environmental conditions, suggesting a potential regulatory function. At least three

sRNAs were found to affect the regulation of RpoS translation (Wassarman et al. 2001). DsrA and RprA stimulate RpoS translation in response to low temperature and cell surface stress, respectively, whereas OxyS represses RpoS translation in response to oxidative shock. However, in addition to regulating RpoS translation, DsrA represses the translation of HNS (a global regulator of gene expression), whereas OxyS represses the translation of FhlA (a transcriptional activator), allowing the cell to coordinate different pathways involved in cell adaptation. The binding of the small noncoding RNAs to its target mRNA is facilitated by a small protein Hfq, which thus plays a critical role in the regulation of expression of the general stress response (Gottesman et al. 2006; Sittka et al. 2008; Vogel and Luisi 2011; Zhang et al. 2003).

Control of the Heat-Shock Response and the General Stress Response in Gram-Positive Bacteria Although the stress gene and proteins in Gram-negative and Gram-positive bacteria are highly conserved, regulation of these genes is very variable. The presence of HrcA-CIRCE control elements has been noted in only some Gram-negative bacteria, and a comparison of Gram-negative with Gram-positive bacteria reveals major differences. > Table 16.2 shows the factors affecting regulation of major stress genes in Gram-positive bacteria and in two Gram-negative bacteria belonging to the Alphaproteobacteria and Gammaproteobacteria. The data indicate that the expression of a stress protein can be under the control of different regulons and also show difference in control elements between the various bacteria. Many of the genes that in Gram-negative bacteria belong to the heat-shock regulon (as their expression is controlled by the heatshock transcriptional activator s32) constitute part of the general

. Table 16.2 Factors affecting regulation of major stress genes. Regulation of major stress genes Gene

Function of gene product Bacteria

Regulon

Transcription during stress

Control element

dnaK

Chaperone

Heat shock

sΑ (s70)

CIRCE

groEL

Chaperone

Gram positive a-proteobacteria

Heat shock

s32

–

g-proteobacteria

Heat shock

s32

–

Gram positive

Heat shock

s70

CIRCE

Stability of gene product

a-proteobacteria

Heat shock

s32

CIRCE

g-proteobacteria

Heat shock

s32

–

Heat shock

s32

Stable Unstable

rpoH

Activator—s32

a-proteobacteria g-proteobacteria

Heat shock

s32, sE

lon, clpP

Proteases

Gram negative

Heat shock

s32

Gram positive

General stress

sB

Abbreviations: CIRCE a conserved inverted repeat control element. There should be no ND in the table, because this is an irrelevant term here. Please leave the table the way I had it, see above. The way you edited here is confusing and even misleading

Bacterial Stress Response

stress response in Gram-positive bacteria. The only genes that are truly ‘‘heat-shock genes’’ in Gram-positive bacteria are the genes coding for the major chaperones—Hsp10 and Hsp60 (GroES and GroEL) and the Hsp70 group (DnaK, DnaJ, and GrpE).

Heat-Shock Response The model organism for studying the heat-shock response in Gram-positive bacteria is B. subtilis. In contrast to E. coli, where most heat-shock proteins are exclusively under the control of the alternative sigma factor s32, Gram-positive bacteria have no heat-shock-specific sigma factor. Rather, the heat-shock response of these bacteria involves the induction of the major chaperones, which is regulated by the HrcA-CIRCE control elements (Hecker and Volker 1990, 1998; Zuber and Schumann 1994) and several groups of proteins regulated by specific control elements, all of which are discussed below. Another major difference is that some of the proteins, which are part of the heat-shock regulon in E. coli (such as the Clp proteases), are part of general stress proteins (GSPs) in B. subtilis, whose induction is regulated by the alternative sigma factor sB. HRCA-CIRCE-Controlled Genes. This system, consisting of the HrcA repressor which binds to the CIRCE inverted repeat, was already described in the section on > ‘‘Heat-Shock Response’’ in Gram-negative bacteria. Though in Gram-negative bacteria this system controls only the groESL operon, its role in Grampositive bacteria is much more central. In the Gram-positive bacteria, the genes coding for Hsp70 (DnaK) and the proteins functionally associated with it are also under the control of HrcA-CIRCE. Thus, this control element regulates the expression of the genes coding for all the major chaperones. Notably, in these bacteria, the genes coding for the group of Hsp70 chaperones are usually organized in one operon: grpE-dnaK-dnaJ. In the group of low G+C Gram-positive bacteria, such as B. subtilis, this operon also contains the gene coding for the HrcA repressor and is hrcA-grpE-dnaK-dnaJ (Segal and Ron 1996a, b). In B. subtilis, the operons regulated by the HrcA-CIRCE system (groESL and dnaK operons) are always transcribed during heat shock by the vegetative sigma factor sA (Yuan and Wong 1995b). This situation is different from the Gram-negative bacteria, in which all the heat-shock operons, including the groESL operon (which contains the CIRCE element), are transcribed by the specific heat shock s32. Recently, GroE itself has been shown to autogenously regulate the transcription of the groE and dnaK operons by the finding that the GroE chaperonin machine modulates the activity of the HrcA repressor (Mogk et al. 1997). Genes Controlled by Additional Repressors. In Streptomyces coelicolor and Streptomyces albus, the groESL1 operon and the groEL2 gene are regulated by tandem CIRCE elements, whereas the dnaK operon encodes its own autoregulatory repressor (Bucca et al. 1995, 1997). Heat-inducible transcription of the dnaK operon (dnaK, grpE, dnaJ, and hspR) initiates from the vegetative promoter. Disruption of hspR led to high and constitutive transcription levels of the dnaK operon but had no effect on the groE expression level (Bucca et al. 1997). Similar to the

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GroE modulation of HrcA activity, DnaK protein forms a specific ATP-independent complex with the Streptomyces HspR repressor, and this interaction is necessary for HspR to bind a dnaKp fragment in gel shift assays (Bucca et al. 2000). The dnaK heat induction model suggested by Bucca et al. suggests a decrease in the availability of DnaK because of the accumulation of heat-damaged proteins (Bucca et al. 2000). This model has many similarities to the heat induction of the s32-dependent transcription in E. coli, a model that will be discussed in detail below. Another heat-shock control element found in S. albus is the RheA, which represses the transcription of hsp18 (encoding a small heat-shock protein) by binding specifically to the hsp18 promoter (Servant and Mazodier 1996; Servant et al. 1999). Transcription analysis of rheA in the S. albus wild type and in rheA mutant strains suggested that RheA represses transcription not only of hsp18 but also of rheA itself (Servant et al. 1999).

The General Stress Response Sigma B-Controlled Genes. Sigma B was found to control a stress-starvation regulon that comprises a very large set of general stress genes (for reviews, see (Hecker et al. 1996; Hecker and Volker 1998)). These sB-dependent genes are strongly induced by heat, ethanol, acid, or salt stress, as well as by starvation for a carbon source, phosphate, and oxygen (Bernhardt et al. 1997, 1999; Buttner et al. 2001; Hecker and Volker 1998). Recent experiments (Petersohn et al. 2001) using gene arrays containing all currently known open reading frames of B. subtilis suggest that as many as 125 genes are under the control of sB. At least 24 of these also seem to be subject to a second, sB-independent stress induction mechanism. Most of the sB-dependent general stress proteins are probably located in the cytoplasm, but 25 contain at least one membrane-spanning domain, and at least six proteins appear to be secreted. This very large stress regulon seems to give a basal level of protection against a large variety of stress conditions. Two groups of signals were found to trigger the induction of sigB, the gene that codes for sB. The first group contains extracellular signals (i.e., glucose, oxygen, or phosphate but not amino acid starvation) that result in a drop of the ATP level (Maul et al. 1995). (Amino acids trigger the induction of ppGpp and keep the ATP pool constant.) The second group of stimuli includes physical stress factors such as heat, salt, and acid stress but not oxidative stress (Hecker and Volker 1998). This group of stimuli induces the synthesis of sB via a twocomponent system (RsbS and RsbT) that changes the balance of a complex network of anti-sigma (RsbW) factor and its agonist (nonphosphorylated RsbV) to activate sB (Akbar and Price 1996; Yang et al. 1996). For the expression of some genes, the involvement of sB is essential, whereas for others it seems to be nonessential because it can be replaced by alternative stress induction mechanisms (Hecker and Volker 1998). Not much is known about many of the 125 GSP genes (Petersohn et al. 2001), and their physiological role in the

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complex general stress response is not understood. The identified GSPs can be assigned to five main groups (Hecker and Volker 1998): (1) Group 1 is the sB-dependent genes that encode subunits of stress-inducible proteases. ClpP, ClpC, and ClpX are probably essential for the renaturation or degradation of misfolded or denatured proteins that accumulate in the cell upon exposure to stress conditions (Gerth et al. 1998; Gottesman 1989). Null mutants of clpC, clpP, and clpX are extremely sensitive to heat, salt, or ethanol stress and much more sensitive than mutants of sigB (Gerth et al. 1998; Kruger et al. 1994; Msadek et al. 1994). (2) Group 2 is the sB-dependent genes that encode general oxidative stress-protective proteins (such as katE, which encodes catalase; (Engelmann et al. 1995)) and the DNA-protecting protein Dps (Antelmann et al. 1997b). Other sB-dependent proteins (such as thioredoxin ClpC, ClpP, and the fifth and sixth gene products of the clpC operon (sms and yacK); (Kaan et al. 1999) may also be involved in adaptation to oxidative stress (Hecker and Volker 1998). (3) The third group is proteins with a putative role in the adaptation to salt or water stress. A proline uptake system encoded by a functional copy of opuE is required by B. subtilis for the use of external proline as an osmoprotectant (Hecker and Volker 1998). However, the physiological role of sB in the expression of opuE is still unclear because exogenously provided proline was used as an osmoprotectant in a sigB mutant (von Blohn et al. 1997). YtxH and GsiB are homologous to plant desiccation proteins, which are involved in water stress protection, and YkzA is a homolog of the E. coli OsmC, which is involved in osmo-adaptation (Maul et al. 1995; Mueller et al. 1992; Varon et al. 1996; Volker et al. 1994). (4) Group 4 is a heterogeneous group of proteins: their role in adaptation to stress is yet to be determined. One of these proteins, GspA (Antelmann et al. 1995), is also induced upon amino acids starvation (Eymann and Hecker 2001) and seems to be involved in the expression of hag, which encodes flagellin, or UDPglucose pyrophosphorylase, which participates in cell wall metabolism (Varon et al. 1993). Some proteins seem to participate in nicotinamide adenine dinucleotide (NAD) synthesis (e.g., nadC and nadE gene products) or might catalyze reduced NAD phosphate (NADP[H])-dependent reactions (Antelmann et al. 1997a, b; Hecker and Volker 1998; Scharf et al. 1998). (Andersson et al. 1998) The fifth group consists of a large number of proteins that, so far, show no significant similarity to known proteins (Petersohn et al. 2001). Several of the general stress operons in Gram-positive bacteria were found to be regulated by more than one control element. The B. subtilis clpC, clpP, and trxA operons are under the control of the vegetative sigma factor sA and the stress sigma factor sB (Gerth et al. 1998; Kruger et al. 1996; Scharf et al. 1998). Although both promoters were used under a number of stress conditions, the induction pattern of the genes varied for the different genes and the particular stress condition. A cis element that contains a heptameric tandem consensus sequence was found upstream of the clpC, clpE, and clpP B. subtilis operons and was shown to be the binding site of the CtsR

repressor (Derre et al. 1999a, b; Kruger and Hecker 1998). CstR was lately found also in Listeria monocytogenes (Nair et al. 2000).

Complexity of the Stress Response Networks Regulation of bacterial stress response involves various positive and negative control elements, which often interact with each other. Some heat-shock proteins are directly regulated by only one control element, but other genes and operons are regulated by several control elements (e.g., E. coli pspABCE (Jovanovic et al. 1996), A. tumefaciens groESL (Nakahigashi et al. 1999; Segal and Ron 1995b, 1996a), and B. subtilis clpC (Gerth et al. 1998; Kruger et al. 1996; Scharf et al. 1998). However, the stress response is always a complex response that regulates itself. As an example, the heat-shock response is induced by damaged proteins, whose cellular concentration increases with temperature. Yet, since the heat-shock stimulon contains the genes coding for proteases and chaperones, their induction at increased temperatures reduces the concentration of the damage proteins, thus reducing the level of induction of the heat-shock response. Because the regulatory elements of these complex stress response networks are associated with each other, any impairment of the cellular steady state at one point may affect the whole network, directly or indirectly. Therefore, the study of these global regulatory networks requires global analysis methods (Rosen and Ron 2002). Such methods for transcriptome and proteome analysis are now available and have been implemented in this field. For comprehensive understanding, more than one method should be used. Analysis of mRNA levels is required to define all the genes whose transcription is affected by environmental conditions or regulatory genes. This analysis, however, is insufficient because the expression and activity of the stress genes are controlled at posttranscriptional, higher regulatory levels. Thus, global analysis at the protein level (i.e., proteomics studies) also must be performed. These studies define the final cellular level of the various proteins, as well as their modifications, some of which may be controlled by stress conditions. One important protein modification shown to play a role in global regulatory networks is protein phosphorylation, usually at one or a few amino acids. Recently, a new group of highly phosphorylated proteins has been identified. These proteins accumulated during several stress conditions and may be involved in the degradation process (Rosen et al. 2004). In eukaryotic systems, protein phosphorylations are known to be involved in protein labeling and in many signal transduction pathways. In bacteria, the number of known phosphorylated proteins is much lower. However, several phosphorylated proteins are involved in the heat-shock response of various bacteria, as will be shown in the following examples. The heat-shock transcriptional activation of the lonD gene of Myxococcus xanthus is controlled by a two-component histidine-aspartate phosphorylation system (Ueki and Inouye 2002). The general stress

Bacterial Stress Response

sigma factor of B. subtilis (sB) is regulated by several regulatory kinases and phosphatases (the Rsb proteins), which catalyze the release of sB from an anti-sB factor (Akbar et al. 2001; Akbar and Price 1996; Yang et al. 1996; Zhang et al. 2001). Another heatshock protein (Hsp70 of Mycobacterium leprae) was found to be phosphorylated at threonine-175 (Peake et al. 1998), which results in an increased affinity for a model polypeptide substrate. One of the best-studied examples of stress-controlled protein modification was already discussed above in the general stress response of E. coli. The phosphorylated form of RssB (a stationary phase response regulator) targets the alternative transcriptional activator sS for degradation by ClpXP (Bouche et al. 1998; Zhou et al. 2001). In view of these examples, protein modification will probably be demonstrated as one of the important control elements in global regulatory networks.

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Shenhar Y, Rasouly A, Biran D, Ron EZ (2009) Adaptation of Escherichia coli to elevated temperatures involves a change in stability of heat shock gene transcripts. Environ Microbiol 11:2989–2997 Sherman M, Goldberg AL (1992) Involvement of the chaperonin dnaK in the rapid degradation of a mutant protein in Escherichia coli. EMBO J 11:71–77 Sherman MY, Goldberg AL (1996) Involvement of molecular chaperones in intracellular protein breakdown. EXS 77:57–78 Singh SS, Typas A, Hengge R, Grainger DC (2011) Escherichia coli sigma senses sequence and conformation of the promoter spacer region. Nucleic Acids Res 39:5109–5118 Sittka A, Lucchini S, Papenfort K, Sharma CM, Rolle K, Binnewies TT, Hinton JC, Vogel J (2008) Deep sequencing analysis of small noncoding RNA and mRNA targets of the global post-transcriptional regulator. Hfq PLoS Genet 4:e1000163 Sparrer H, Rutkat K, Buchner J (1997) Catalysis of protein folding by symmetric chaperone complexes. Proc Natl Acad Sci USA 94:1096–1100 Srivastava P (2002) Roles of heat-shock proteins in innate and adaptive immunity. Nature Rev Immunol 2:185–194 Storz G, Vogel J, Wassarman KM (2011) Regulation by small RNAs in bacteria: expanding frontiers. Mol Cell 43:880–891 Strauch KL, Johnson K, Beckwith J (1989) Characterization of degP, a gene required for proteolysis in the cell envelope and essential for growth of Escherichia coli at high temperature. J Bacteriol 171:2689–2696 Straus DB, Walter WA, Gross CA (1987) The heat shock response of E. coli is regulated by changes in the concentration of sigma 32. Nature 329:348–351 Straus DB, Walter WA, Gross CA (1989) The activity of sigma 32 is reduced under conditions of excess heat shock protein production in Escherichia coli. Genes Dev 3:2003–2010 Straus D, Walter W, Gross CA (1990) DnaK, DnaJ, and GrpE heat shock proteins negatively regulate heat shock gene expression by controlling the synthesis and stability of sigma 32. Genes Dev 4:2202–2209 Studemann A, Noirclerc-Savoye M, Klauck E, Becker G, Schneider D, Hengge R (2003) Sequential recognition of two distinct sites in sigma(S) by the proteolytic targeting factor RssB and ClpX. EMBO J 22:4111–4120 Taura T, Kusukawa N, Yura T, Ito K (1989) Transient shut off of Escherichia coli heat shock protein synthesis upon temperature shift down. Biochem Biophys Res Commun 163:438–443 Tilly K, McKittrick N, Zylicz M, Georgopoulos C (1983) The dnaK protein modulates the heat shock response of Escherichia coli. Cell 34:641–646 Tilly K, Spence J, Georgopoulos C (1989) Modulation of stability of the Escherichia coli heat shock regulatory factor sigma. J Bacteriol 171:1585–1589 Tomoyasu T, Gamer J, Bukau B, Kanemori M, Mori H, Rutman AJ, Oppenheim AB, Yura T, Yamanaka K, Niki H et al (1995) Escherichia coli FtsH is a membrane-bound, ATP-dependent protease which degrades the heat-shock transcription factor sigma 32. EMBO J 14:2551–2560 Tomoyasu T, Mogk A, Langen H, Goloubinoff P, Bukau B (2001) Genetic dissection of the roles of chaperones and proteases in protein folding and degradation in the Escherichia coli cytosol. Mol Microbiol 40:397–413 Typas A, Barembruch C, Possling A, Hengge R (2007a) Stationary phase reorganisation of the Escherichia coli transcription machinery by Crl protein, a fine-tuner of sigmaS activity and levels. EMBO J 26:1569–1578 Typas A, Becker G, Hengge R (2007b) The molecular basis of selective promoter activation by the sigmaS subunit of RNA polymerase. Mol Microbiol 63:1296–1306 Typas A, Stella S, Johnson RC, Hengge R (2007c) The 35 sequence location and the Fis-sigma factor interface determine sigmas selectivity of the proP (P2) promoter in Escherichia coli. Mol Microbiol 63:780–796 Ueki T, Inouye S (2002) Transcriptional activation of a heat-shock gene, lonD, of Myxococcus xanthus by a two component histidine-aspartate phosphorelay system. J Biol Chem 277:6170–6177 Val DL, Cronan JE Jr (1998) In vivo evidence that S-adenosylmethionine and fatty acid synthesis intermediates are the substrates for the LuxI family of autoinducer synthases. J Bacteriol 180:2644–2651 van Asseldonk M, Simons A, Visser H, de Vos WM, Simons G (1993) Cloning, nucleotide sequence, and regulatory analysis of the Lactococcus lactis dnaJ gene. J Bacteriol 175:1637–1644

Bacterial Stress Response Van Bogelen RA, Kelley PM, Neidhardt FC (1987) Differential induction of heat shock, SOS and oxidation stress regulons and accumulation of nucleotides in Escherichia coli. J Bacteriol 169:26–32 Varon D, Boylan SA, Okamoto K, Price CW (1993) Bacillus subtilis gtaB encodes UDP-glucose pyrophosphorylase and is controlled by stationary-phase transcription factor sigma B. J Bacteriol 175:3964–3971 Varon D, Brody MS, Price CW (1996) Bacillus subtilis operon under the dual control of the general stress transcription factor sigma B and the sporulation transcription factor sigma H. Mol Microbiol 20:339–350 Vicente M, Chater KF, De Lorenzo V (1999) Bacterial transcription factors involved in global regulation. Mol Microbiol 33:8–17 Vogel J, Luisi BF (2011) Hfq and its constellation of RNA. Nat Rev Microbiol 9:578–589 Vogel J, Bartels V, Tang TH, Churakov G, Slagter-Jager JG, Huttenhofer A, Wagner EG (2003) RNomics in Escherichia coli detects new sRNA species and indicates parallel transcriptional output in bacteria. Nucleic Acids Res 31:6435–6443 Volker U, Engelmann S, Maul B, Riethdorf S, Volker A, Schmid R, Mach H, Hecker M (1994) Analysis of the induction of general stress proteins of Bacillus subtilis. Microbiology 140(Pt 4):741–752 von Blohn C, Kempf B, Kappes RM, Bremer E (1997) Osmostress response in Bacillus subtilis: characterization of a proline uptake system (OpuE) regulated by high osmolarity and the alternative transcription factor sigma B. Mol Microbiol 25:175–187 Waldminghaus T, Gaubig LC, Klinkert B, Narberhaus F (2009) The Escherichia coli ibpA thermometer is comprised of stable and unstable structural elements. RNA Biol 6:455–463 Wang QP, Kaguni JM (1989) dnaA protein regulates transcriptions of the rpoH gene of Escherichia coli. J Biol Chem 264:7338–7344 Wassarman KM, Storz G (2000) 6S RNA regulates E. coli RNA polymerase activity. Cell 101:613–623 Wassarman KM, Repoila F, Rosenow C, Storz G, Gottesman S (2001) Identification of novel small RNAs using comparative genomics and microarrays. Genes Dev 15:1637–1651 Wawrzynow A, Wojtkowiak D, Marszalek J, Banecki B, Jonsen M, Graves B, Georgopoulos C, Zylicz M (1995) The ClpX heat-shock protein of Escherichia coli, the ATP-dependent substrate specificity component of the ClpP-ClpX protease, is a novel molecular chaperone. EMBO J 14:1867–1877

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17 Anaerobic Biodegradation of Hydrocarbons Including Methane Johann Heider . Karola Schu¨hle Fachbereich Biologie, Laboratorium fu¨r Mikrobiologie, Marburg, Germany

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 605 Substrates and Ecology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 606 Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 606 Contaminated Sites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 608 Natural Gas and Oil Seeps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 608 Deep Subsurface Environments and Oil Reservoirs . . . . . 610 Growth with Crude Oil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 610 Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 610 Denitrifying and Nitrate-Ammonifying Bacteria . . . . . . . 611 Metal Ion Reducers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 612 Sulfate-Reducing Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . 612 Phototrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 612 Methanogenic Consortia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 613 Sulfidogenic Methane-Degrading Consortia . . . . . . . . . . . . 614 Fermentative Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 614 ‘‘Intra-aerobic’’ Anaerobes Deriving O2 from Either Chlorate or Nitrite . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 614 Cultivation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 615 Cultivation Methods with Hydrocarbons . . . . . . . . . . . . . . . 617 Biochemistry of Microbial Hydrocarbon Degradation . . . . 617 Fumarate Addition Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . 618 Oxygen-Independent Hydroxylation . . . . . . . . . . . . . . . . . . . . 622 Carboxylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 624 Acetylene Hydration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 624 Alkene Hydration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 625 ‘‘Reverse Methanogenesis’’ by Sulfate-Reducing Consortia . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 626 Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 627 Bioremediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 627 Microbial Enhanced Oil Recovery and Biofuels . . . . . . . . . 628 Fine Chemicals/Biotechnology . . . . . . . . . . . . . . . . . . . . . . . . . . 628 Abstract Hydrocarbons are highly abundant in nature, originating from both, geochemical and biological processes. Because of the low reactivity of their C-H bonds, either aliphatic or aromatic, hydrocarbons are usually hard to degrade biologically, and all known aerobic hydrocarbon-degrading organisms overcome this reactivity barrier by involving oxygen-dependent monoor dioxygenases for the initial attack at the hydrocarbons.

It was surprising when about 20 years ago biological hydrocarbon degradation was shown to occur also in the absence of molecular oxygen. In the meantime, many hydrocarbons are known to be degraded anaerobically by a large variety of facultatively or strictly anaerobic bacteria. These bacteria initiate their metabolic pathways by oxygen-independent hydrocarbon-activating reactions that are completely different and much more divergent than the “aerobic” initial reactions and represent novel biochemical principles. We review here the current knowledge on the ecology of anaerobic hydrocarbon degraders and their habitats, introduce the organisms involved, and describe the biochemical pathways and enzymes allowing these remarkable properties. Finally, we discuss some implications of these new insights on possible applications.

Introduction Hydrocarbons are all chemical compounds containing only carbon and hydrogen atoms. The lack of polar functional groups causes a generally low solubility of these compounds in water, and the high stability of the C–H bonds results in low chemical reactivity of most hydrocarbons (Wilkes and Schwarzbauer 2010). They are arranged into two main groups: the aliphatic (inclusively alicyclic) and the aromatic hydrocarbons. The most abundant among the aliphatic hydrocarbons are the alkanes, which contain exclusively saturated bonds and are highly stable (hence the alternative name ‘‘paraffins’’). In addition, a number of unsaturated hydrocarbons are known to occur naturally, either alkenes containing one or more double bonds or alkynes containing triple bonds. Both alkenes and alkynes are significantly more reactive than the corresponding alkanes, due to the energyrich status of the respective C–C multiple bonds. The alicyclic hydrocarbons are a separate class of aliphatic compounds containing nonaromatic ring structures. The second main group embraces the aromatic hydrocarbons, which also occur in vast amounts. They always contain one or more aromatic rings, which may carry additional side chains. The aromatic rings may be fused to highly complex structures. The low reactivity of the aromatic hydrocarbons compared to alkenes can be explained by the aromatic stabilization effect, which makes it catalytically highly demanding to attack the rings chemically. Biological degradation of any hydrocarbon substrates requires an initial introduction of a functional group where further biochemical reactions can occur. Therefore, the initial reactions attacking hydrocarbons are of special biochemical interest.

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_80, # Springer-Verlag Berlin Heidelberg 2013

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Hydrocarbons are widespread in natural environments and are naturally formed either by slow geochemical processes (Tissot and Welte 1984) or as normal metabolic products of many microorganisms, plants, or animals (Birch and Bachofen 1988; Widdel and Rabus 2001). Due to human use of petroleum products, the environmental exposure to hydrocarbons has been increased considerably. It is therefore no big surprise that many microorganisms are able to degrade all kinds of hydrocarbons, and the first examples of aerobic hydrocarbon-degrading bacteria or fungi have been known since the beginning of the twentieth century. The microbiology and biochemistry involved in the aerobic metabolism of hydrocarbons in these organisms are known in detail (see > Chap. 5, ‘‘Hydrocarbon-Oxidizing Bacteria’’ in this volume), and in every case the initial reaction of any aerobic hydrocarbon degrader is catalyzed by an oxygendependent enzyme, either a monooxygenase or a dioxygenase (Widdel and Musat 2010). These enzymes create a highly potent oxidizing agent from oxygen in their active site, which overcomes the low reactivity of the hydrocarbons and allows to introduce functional groups (e.g., hydroxyl groups) that enables further biochemical conversions. Therefore, it was long regarded as impossible to degrade hydrocarbons in the absence of oxygen, and it took until the 1990s to conclusively prove the existence of hydrocarbon degradation also under anoxic conditions. The organisms responsible for anaerobic hydrocarbon degradation belong to different taxonomic groups of anaerobic or facultatively anaerobic bacteria and Archaea. Most of these organisms use anaerobic respiration with nitrate, metal ions, or sulfate for their energy metabolism, but there are also examples of anoxygenic phototrophs, fermentative or syntrophic microorganisms known to degrade hydrocarbon substrates (Heinnickel et al. 2010; Widdel and Grundmann 2010; Widdel and Musat 2010). The biochemical mechanisms employed in anaerobic hydrocarbon metabolism are much more diverse than the oxygen-dependent hydroxylation reactions used in aerobic hydrocarbon metabolism (Widdel and Musat 2010). There are examples for at least six different principles of oxygen-independent hydrocarbon activation used in different organisms and/or for the degradation of different hydrocarbons, which are (1) radical-based addition of a fumarate cosubstrate to methyl or methylene groups of hydrocarbons, (2) oxygenindependent hydroxylation of secondary or tertiary C-atoms, (3) carboxylation of unsubstituted aromatic rings, (4) hydration of the triple bond of acetylene, (5) hydration of the double bonds of alkenes, and (6) activation of methane by ‘‘reverse methanogenesis’’ (Harder 2010; Heider and Boll 2010; Taupp et al. 2010; Tierney and Young 2010). Finally, it has recently been demonstrated that some ‘‘intra-aerobic’’ microorganisms can generate molecular oxygen from chlorite or nitrite and use it for mono- or dioxygenase-based functionalization of hydrocarbons even in anoxic environments (Mehboob et al. 2010). Most of the known anaerobic hydrocarbon modifications are slower than the corresponding aerobic reactions, and the enzymes involved are usually highly oxygen-sensitive. Therefore, anaerobic hydrocarbon degraders do not usually degrade these substrates under oxic conditions,

unless they can shift from the anaerobic pathway to an aerobic oxygenase-based pathway, which was only found for very few organisms (Shinoda et al. 2004). This chapter presents the current knowledge on the microbiology, ecology, and biochemistry of anaerobic hydrocarbon metabolism. We first introduce the types of hydrocarbons that are degraded anaerobically and the environments and ecological niches where the respective organisms are found. The microorganisms involved in the process are then introduced in detail, followed by protocols for their enrichment and cultivation. Moreover, the biochemical pathways and the enzymes involved in catalyzing the key steps of the known oxygen-independent hydrocarbon degradation strategies are presented. Finally, some possible applications of this metabolic capacity of microorganisms are discussed. Further information on anaerobic hydrocarbon degradation can be obtained from many review articles, of which we recommend a few very recent examples here: for a general overview on the topic, we refer the reader to Heider (2007); Foght (2008); Carmona et al. (2009); Rabus et al. (2009); Heider and Boll (2010); Widdel and Grundmann (2010); Widdel and Musat (2010); for more specialized summaries on anaerobic methane oxidation, see Knittel and Boetius (2009); Gray et al. (2010); Taupp et al. (2010); for anaerobic oxidation of benzene and other more recalcitrant hydrocarbons, see Meckenstock and Mouttaki (2011); Vogt et al. (2011); and for reviews on applied aspects, see Brown (2010); Gray et al. (2010).

Substrates and Ecology Substrates Hydrocarbons known to be degraded anaerobically belong to all structural classes, alkanes, alkenes, alkynes, alicyclic and aromatic hydrocarbons, as shown in > Fig. 17.1. Among the alkanes, the chemically simplest hydrocarbon methane as well as higher n-alkanes from propane to chain lengths of more than 20 C-atoms are included (Aeckersberg et al. 1991; Ehrenreich et al. 2000; Kniemeyer et al. 2007; Widdel and Grundmann 2010). No reliable data are available on any anaerobic metabolic conversion of ethane, which has only recently been shown to be degraded aerobically by newly discovered bacteria at marine hydrocarbon seeps, possibly involving a specialized methane monooxygenase-like enzyme (Redmond et al. 2010). Branched alkanes are known to be degraded under anoxic conditions (Bregnard et al. 1997), although the organisms and pathways involved are unknown. Regarding anaerobic degradation of alicyclic alkanes, a highly enriched cyclohexane-degrading culture has recently been described, which apparently couples nitrate reduction with the removal of toxic nitrite via anaerobic ammonium oxidation (Musat et al. 2010). Straight-chain 1-alkenes are readily degraded anaerobically, often by the same organisms capable of anaerobic metabolism of n-alkanes of similar chain length. Another class of alkenes known to be degraded in anoxic environments are terpenoids (Harder 2010).

Anaerobic Biodegradation of Hydrocarbons Including Methane

H H

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Propane

C

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. Fig. 17.1 Hydrocarbon substrates known to be degraded anaerobically

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. Fig. 17.2 Free Gibbs energies for anaerobic oxidation of alkanes (a) and 1-alkenes (b) of increasing lengths combined with sulfate reduction. The calculated overall DG ’ values of the reactions (circles, left ordinates) as well as the relative DG ’ values per carbon atom of the hydrocarbons (squares, right ordinates) are given

Comparison of the thermodynamics of alkane and alkene degradation under sulfate-reducing conditions reveals that the C1 and C2 compounds must behave quite differently from those with larger chain lengths: among the alkanes, methane oxidation yields much less energy per carbon atom ( 13.1 kJ/mol) than that of the larger alkanes, which level off at values of 31 to 33 kJ/mol (per C-atom) for propane and higher alkanes (> Fig. 17.2). Among the 1-alkenes, the (theoretical) oxidation of ethylene coupled to sulfate reduction would yield 60 kJ/mol per C-atom, whereas the calculated energy yields decrease with increasing chain length, until they reach values of 36 to 40 kJ/mol per C-atom (> Fig. 17.2). Despite the apparently favorable thermodynamics, no data are available on the anaerobic degradability of ethylene or other short-chain alkenes,

nor have any long-chain 2- or 3-alkenes been tested. Among the alkynes, only acetylene is known to be degraded anaerobically via a very special hydration reaction to acetaldehyde (Ro¨sner and Schink 1995). Finally, many aromatic hydrocarbons are degraded under anoxic conditions, but the substrate range seems to be restricted to monocyclic, bicyclic, and very few tricyclic compounds. The best biodegradability is found for alkyl-substituted mono- or bicyclic hydrocarbons, such as toluene, ethylbenzene, the xylenes or methylnaphthalene, whereas the non-substituted aromatic ring systems, such as benzene or naphthalene, are only very poor substrates for anaerobic degradation. The only tricyclic compounds known to be anaerobically degraded are phenanthrene and anthracene (Tierney and Young 2010), whereas most other highly

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Anaerobic Biodegradation of Hydrocarbons Including Methane

condensed aromatic systems seem to be inert in anoxic environments. These types of hydrocarbons are apparently only degradable with the help of molecular oxygen. The occurrence of anaerobic biological degradation of the hydrocarbons listed above is firmly documented by the isolation of pure cultures of bacterial strains growing on one or more defined hydrocarbon substrates. Moreover, fundamentally different metabolic pathways are used for anaerobic degradation of some model hydrocarbons, compared to those used by aerobic hydrocarbon-degrading organisms. The important role of anaerobic hydrocarbon degradation in natural environments is confirmed by studying the on-site prevalence of characteristic microorganisms or metabolic intermediates, as well as the isotopic signatures of the hydrocarbons and the derived metabolites, which are in accord with the expected biological processes (Meckenstock et al. 2004).

Contaminated Sites Anaerobic hydrocarbon degradation is routinely observed in anoxic environments contaminated with petroleum or tar by-products, which are regularly and inadvertently created by industrial processing and handling of products from petroleum and coal by humans. Therefore, extensive hydrocarbon contamination is often found at gas stations or factory sites, e.g., chemical industry, petroleum refineries or old gasworks, and coke ovens (Rios-Hernandez et al. 2003; Meckenstock et al. 2004; Vieth et al. 2005; Parisi et al. 2009; Jobelius et al. 2011). While aerobic hydrocarbon degraders dominate at the surfaces exposed to air, oxygen soon becomes limiting when the hydrocarbon contaminants permeate deeper into soil or sediment layers, because microbial metabolic activity leads to anaerobiosis already within millimeters of a waterlogged sediment. Of special interest are contaminations in groundwater aquifers, which lead to the development of pollutant plumes that may impair large surrounding areas, endanger the drinking water supply, or threaten to contaminate rivers. During the last two decades, it has been established that these contaminated environments are in fact excellent habitats for all kinds of anaerobic hydrocarbon-degrading bacteria. The occurrence of different physiological groups of these organisms is correlated to the chemical composition of the water, especially with regard to the available electron acceptors for anaerobic respiration (Beller et al. 2002; Kunapuli et al. 2007; Winderl et al. 2010; Kuntze et al. 2011). In most cases, gradients are observed over the extent of the pollutant plumes, which exhibit aerobic hydrocarbon degradation at the edges, where the contaminant concentrations are low enough to be degraded with the little oxygen present in groundwater, followed by anaerobic hydrocarbon degradation by a succession of different physiological groups of anaerobic respiratory bacteria. The observed succession is governed by the available energy from using the respective electron acceptors, beginning with denitrifying bacteria, followed by metal-ion- (e.g., FeIII or MnIV) and sulfate-reducing bacteria, and ending with methanogenic

consortia of hydrocarbon-metabolizing bacteria and methanogenic Archaea. The extent of these zones is mainly dependent on the relative amounts of the respective electron acceptors at the individual sites. However, the abundance and identity of the correlated bacteria must also be affected by additional still unknown factors because very different bacterial communities may be present in contaminated aquifers where degradation follows the same physiological regime (e.g., FeIII-reducing sites that may either be dominated by Geobacter- or by Georgfuchsia-type bacteria) (Weelink et al. 2009; Pilloni et al. 2011).

Natural Gas and Oil Seeps Other important habitats of anaerobic hydrocarbon-degrading microorganisms are natural gas and oil seeps. These are sites of leakage of natural oil and gas deposits, which may be localized either on land or on the sea floor. The most abundant reservoir of stored hydrocarbons on earth consists of huge deposits of methane hydrate in deep sea or subsurface permafrost environments all around the continental shelves, which are estimated to exceed the total amount of natural gas from other sources two- to tenfold (> Fig. 17.3). These structures are formed from methane of either geochemical or microbial origin at high pressure and low temperatures and represent clathrate compounds of the hydrocarbon molecules encased in cages of water molecules (Crabtree 1995; Kvenvolden 1999). Slow decomposition of methane hydrates is believed to drive marine gas seeps that have been detected at many places on the continental shelves and found to provide energy for the development of rich deep-sea communities of specialized animals that basically depend on chemotrophic methanedegrading bacteria. Whereas most of these bacteria represent aerobic methane degraders, geochemical studies in marine sediments suggested the existence of anaerobic methane oxidation already 40 years ago (Martens and Berner 1974; Barnes and Goldberg 1976; Reeburgh 1976). These studies were corroborated by in situ analysis of metabolite patterns in the sediments, 14C-methane injection experiments, and 13 C-discrimination data (Reeburgh 1980; Alperin and Reeburgh 1985; Iversen and Jorgensen 1985; Ritger et al. 1987; Paull et al. 1992; Hansen et al. 1998). Final proof came with the detection of the first true anaerobic methane-degrading microbial consortia, which are directly associated with deep-sea methane hydrate deposits in anoxic sediments (Boetius et al. 2000). A special case was found at the frequent gas seeps on the floor of the Black Sea, which exhibits totally anoxic conditions already in the water column below 300 m, rather than only in the sediment. Therefore, no aerobic methane degradation occurs there in deeper water layers, and the anaerobic methane-oxidizing prokaryotes on the ground can grow up several meters to build extensive mats and reef-like structures (Michaelis et al. 2002). Likewise, marine petroleum seeps are known that are fueled by larger hydrocarbons leaking out of offshore oil deposits (Wilkinson 1972) (> Fig. 17.4). Similar as described for the

Anaerobic Biodegradation of Hydrocarbons Including Methane

17

. Fig. 17.3 Localization of known methane hydrate deposits

. Fig. 17.4 Localization of some oil and gas seeps off the coast of California. Closed circles: oil seeps, open circles: gas seeps, half-filled circles: tar seeps, stars: oil and gas seeps, open squares: development platforms (After Wilkinson 1972)

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Anaerobic Biodegradation of Hydrocarbons Including Methane

methane-driven gas seeps, these sites are populated by large populations of aerobic and anaerobic hydrocarbon-degrading prokaryotes which provide the basis of thriving deep-sea ecosystems fueled independently of solar light (LaMontagne et al. 2004). Anaerobic methane-oxidizing microbial consortia are also abundantly present at these sites, which suggests that degradation of larger hydrocarbons by methanogenic consortia simultaneously feeds anaerobic methane oxidation (LaMontagne et al. 2004; Gray et al. 2010; Lloyd et al. 2010). Terrestrial seeps are characterized either as petroleum/tar seeps, which often indicate the presence of exploitable oil deposits, or as mud volcanoes, the equivalent of marine gas seeps, which are mostly driven by the decomposition of methane hydrate in the underground and contain large amounts of methane and other gaseous hydrocarbons in their exhaust. Anaerobic hydrocarbon degraders are known from both types of environments (Alain et al. 2006; Gray et al. 2010; Lloyd et al. 2010). It is obvious that the marine sites are usually dominated by sulfate-reducing microbial communities because of the large sulfate concentration of seawater (28 mM), whereas the organisms present at terrestrial sites should be expected to vary more widely in their physiology, dependent on the prevalent electron acceptors in the local water supply.

Deep Subsurface Environments and Oil Reservoirs A relatively new development in the last decade is based on the finding of apparent biodegradation going on in situ even in deep subsurface environments, especially oil reservoirs, as indicated by some initial reports of methane production from alkanes by anaerobic enrichment cultures (Zengler et al. 1999b). The in situ processes take place at elevated but not exceedingly high temperatures found in some oil reservoirs (45–70  C). They seem to be mostly mediated by methanogenic consortia, circumventing the need of replenishing the electron acceptors (Gray et al. 2010), but in some instances also by sulfate-reducing communities (Gieg et al. 2010). Biogenic methane production from crude oil hydrocarbons was shown at many oil-drilling sites (Nazina et al. 2006; Mochimaru et al. 2007; Jones et al. 2008; Gray et al. 2009; Parisi et al. 2009; Gieg et al. 2010), and the connection of this process to the well-known ‘‘weathering’’ of oil deposits has been recognized (Gray et al. 2010). The more the biodegradation continues in an oil reservoir, the more of the easily degradable hydrocarbons will be lost, leading to vast deposits of heavy oil and tar, which are difficult and expensive to exploit. It is even possible to link the amount and the manner of biodegradation both to the isotopic signature of the oil and to the presence of characteristic marker molecules that are formed as intermediates during hydrocarbon metabolism and are getting enriched in the oil reservoir with the degree of biodegradation (Wardlaw et al. 2008; Gray et al. 2010; Yagi et al. 2010). The microbial composition found in production waters of oildrilling operations is very similar to that of other methanogenic

habitats where hydrocarbons are degraded (Nazina et al. 2006; Mochimaru et al. 2007; Gieg et al. 2010; Gray et al. 2010), as are the patterns of preferentially consumed hydrocarbons (Jones et al. 2008; Gray et al. 2010). Therefore, the deep subsurface organisms responsible for oil weathering in the reservoirs seem to be very similar to those found at surface seep sites or in contaminated aquifer systems. Because geochemical transformation processes of organic materials occur in all subsurface environments, microbial hydrocarbon degradation processes are to be expected even at ‘‘normal’’ sites containing only small amounts of finely dispersed carbonaceous material that does not suffice for the formation of oil reservoirs.

Growth with Crude Oil Changes occurring in the composition of crude oil during the growth of anaerobic hydrocarbon-degrading enrichments or pure cultures under either sulfate-reducing or denitrifying regime have been repeatedly investigated (Rueter et al. 1994; Rabus et al. 1996, 1999; Rabus and Widdel 1996; Wilkes et al. 2000, 2003). Regardless of the physiological regime of the experiments, it appears that the easily degradable n-alkanes ranging from chain lengths of C6–C16 are always removed first, along with the easily degradable aromatic compounds toluene, ethylbenzene, and the xylenes. The total fraction of crude oil to be degraded fast was in the range of 10 % (w/w) of the total hydrocarbons. The same observations were recently also reported for naturally occurring oil degradation directly in the reservoirs (Jones et al. 2008; Gray et al. 2010). A quantitative production of methane was found to be coupled to the selective degradation of n-alkanes from crude oil, and comparison of oil reservoirs differently affected by biological weathering showed that the n-alkanes and alkylbenzenes were always removed first, leaving over the heavier and harder degradable compounds (Connan et al. 1996). Metabolism of alkenes and other compounds known to be easily degradable could not be determined in these experiments because of their low contents in crude oil.

Organisms The organisms involved in anaerobic hydrocarbon degradation are mostly anaerobically respiring bacteria. This solves the problem of obtaining enough energy for growth from degrading the relatively energy-poor substrates by coupling their degradation to the respective anaerobic respiratory chains, while there is still no oxygen available for the typical mono- or dioxygenasemediated initial reactions of aerobic hydrocarbon degraders. The type of electron acceptor in these organisms can be virtually anything that works for any anaerobic respiration, ranging from nitrate, nitrite, or nitrous oxide over oxidized metal ions, sulfate and thiosulfate to protons, which are reduced to hydrogen by hydrocarbon degraders present in methanogenic consortia.

Anaerobic Biodegradation of Hydrocarbons Including Methane

Denitrifying and Nitrate-Ammonifying Bacteria Most of the known denitrifying hydrocarbon-degrading bacteria belong to the order Rhodocyclales in the Betaproteobacteria. Strains degrading toluene, ethylbenzene, xylenes, p-cymene, p-ethyltoluene, several monoterpenes, or even n-hexane have been isolated over the last two decades (> Table 17.1). They are affiliated to the genera Thauera, Azoarcus, Aromatoleum, and Georgfuchsia. Within the genus Thauera, the species T. aromatica contains toluene- or m-xylene-degrading strains, whereas the species T. terpenica and T. linaloolentis contain strains degrading terpenes, p-cymene or p-ethyltoluene (summarized in Anders et al. 1995; Foss and Harder 1998; Heider and Fuchs 2005). Many of the other described anaerobic hydrocarbon-degrading isolates were affiliated to the genus Azoarcus, which was first defined for nitrogen-fixing plant symbionts that are not capable of anaerobic degradation of hydrocarbons or aromatic

17

compounds (Reinhold-Hurek and Hurek 2000). Formally described toluene-degrading species are A. toluvorans, A. tolulyticus, and A. toluclasticus, while many further hydrocarbon-degrading related strains have not yet been formally described as species (Zhou et al. 1995; Song et al. 1999; Reinhold-Hurek et al. 2005). Because the anaerobic degraders of aromatic or hydrocarbon substrates and the nitrogen-fixing plant symbionts currently placed in the genus Azoarcus seem to be only distantly related, the former group of strains will be regrouped under the new candidatus genus name Aromatoleum (F. Widdel, R. Rabus, personal communication). Among the most remarkable of these strains are Aromatoleum aromaticum EbN1, the only known isolate growing anaerobically either on toluene or on ethylbenzene and the first to be genomesequenced (Rabus et al. 2005), and strain HxN1, which grows anaerobically on n-hexane and some other n-alkanes (Ehrenreich et al. 2000). Finally, Georgfuchsia is a recently

. Table 17.1 Representative denitrifying hydrocarbon-degrading bacteria. Phylogenetic rank and the range of hydrocarbons degraded are given. For the known strains without valid taxonomic description, the closest validly described relative is indicated (‘‘aff.’’ for affiliated) Organism

Phylogeny

Substrate range

Described in

Betaproteobacteria

Toluene

Anders et al. (1995)

Azoarcus tolulyticus (e.g., strains Tol 14, Td15)

Toluene, m-xylene

Zhou et al. (1995)

Azoarcus toluvorans

Toluene

Song et al. (1999)

(Facultative) anaerobic bacteria Thauera aromatica (e.g., strains K172, T1)

Azoarcus toluclasticus

Toluene

Song et al. (1999)

Azoarcus sp. strain T

Toluene, m-xylene

Reinhold-Hurek et al. (2005)

Aromatoleum aromaticum strain EbN1

Toluene, ethylbenzene

Rabus et al. (2005)

Strain PbN1 (aff. Aromatoleum)

Ethylbenzene, propylbenzene

Reinhold-Hurek et al. (2005)

Strain ToN1 (aff. A. toluvorans)

Toluene

Reinhold-Hurek et al. (2005)

mXyN1 (aff. T. aromatica)

Toluene, m-xylene

Heider and Fuchs (2005)

pCyN1 (aff. Aromatoleum)

Toluene, p-cymene, various alkenoic monoterpenes

Reinhold-Hurek et al. (2005)

Thauera linaloolentis

Various alkenoic monoterpenes

Foss and Harder (1998)

Thauera terpenica

Various alkenoic monoterpenes

Heider and Fuchs (2005)

pCyN2 (aff. T. terpenica)

Toluene, p-cymene, various alkenoic monoterpenes

Heider and Fuchs (2005)

Castellaniella defragrans

Various alkenoic monoterpenes

Foss et al. (1998)

Strain HxN1 (aff. Aromatoleum)

C6–8 alkanes

Ehrenreich et al. (2000)

Strain OcN1 (aff. Aromatoleum)

C8–12 alkanes

Ehrenreich et al. (2000); Zedelius et al. (2011)

‘‘Intra-aerobic’’ bacteria Strain HdN1

Gammaproteobacteria C14–20 alkanes

Ehrenreich et al. (2000); Zedelius et al. (2011)

Methylomirabilis oxyfera

Phylum NC10

Methane

Ettwig et al. (2010)

Dechloromonas aromatica

Betaproteobacteria

Benzene, toluene, ethylbenzene, xylenes Coates et al. (2001)

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Anaerobic Biodegradation of Hydrocarbons Including Methane

described new genus of strictly anaerobic members of the Rhodocyclaceae capable of anaerobic toluene degradation. This genus seems to be highly abundant in contaminated aquifers and is apparently able to shift its physiology coupled to hydrocarbon metabolism between denitrification and Fe(III) reduction (Weelink et al. 2009). A further species of the Rhodocyclaceae was recognized as anaerobic degrader of terpenoid hydrocarbons, namely, the species Thauera terpenica, whereas the related species T. linaloolentis is only known to degrade polar terpenoids (Foss and Harder 1998). At least one of the known strains of T. terpenica uses alkenes such as a-sabinene, a-phellandrene, b-pinene, and a- or g-terpinene as substrates under denitrifying growth conditions (Foss and Harder 1998). The closely affiliated strain pCyN2 has been reported to degrade the aromatic terpenoid hydrocarbon p-cymene (Harms et al. 1999; Heider and Fuchs 2005), whereas another anaerobic p-cymene-degrading strain, pCyN1 (Harms et al. 1999), is closely related to the candidatus genus Aromatoleum, based on the 16S rRNA sequences (> Table 17.1). Another betaproteobacterial species capable of degrading terpenoid hydrocarbons (e.g., myrcene) under denitrifying conditions is Castellaniella defragrans (Foss et al. 1998; Brodkorb et al. 2010). This bacterium belongs to the Alcaligenaceae in the order Burkholderiales, not to the Rhodocyclales, as most other hydrocarbon-degrading Betaproteobacteria. Denitrifying bacteria from other taxonomic groups have only rarely been described as anaerobic hydrocarbon degraders, with the most notable exception of a strain of the alphaproteobacterial genus Magnetospirillum, which grows under denitrifying conditions on toluene or other polar aromatic compounds and uses the same pathway for metabolizing toluene as all other known anaerobic toluene degraders (Shinoda et al. 2005). Finally, nitrate respiration in the form of nitrate ammonification is an alternative physiological regime for the hydrocarbon-degrading strains of the Fe(III)-reducing Geobacter species (Ahrendt et al. 2007), under which these bacteria can equally well metabolize toluene and other aromatic substrates (Kung et al. 2009).

Metal Ion Reducers The most important described genera representing metal-ionreducing anaerobic hydrocarbon degraders are the strictly anaerobic Geobacter (Deltaproteobacteria) (Lovley et al. 1989; Lovley and Lonergan 1990) and Georgfuchsia (Betaproteobacteria) (Weelink et al. 2009). So far, toluene is the only hydrocarbon substrate known to be degraded by pure cultures of metal-ion-reducing strains, but it is to be expected from field studies that the overall degradation potential of this group of organisms is comparable to that of the denitrifying or sulfate-reducing strains (Weelink et al. 2009; Pilloni et al. 2011; Staats et al. 2011). The most common electron acceptor used is Fe(III), but also other oxidized metal compounds containing,

e.g., Mn(IV) or even U(VI) are used as electron acceptors by some strains (Shelobolina et al. 2008; Weelink et al. 2009). Interestingly, both species are able to shift over to nitrate reduction: denitrification in case of Georgfuchsia (Weelink et al. 2009) and nitrate ammonification in case of Geobacter (Ahrendt et al. 2007). Genome sequences are available of several Geobacter species capable of anaerobic toluene metabolism.

Sulfate-Reducing Organisms Sulfate-reducing bacteria and Archaea are well represented among the known hydrocarbon-degrading isolates or enrichments. They can degrade either alkylbenzenes like toluene, xylenes, or ethylbenzene, long- and short-chain alkanes and 1-alkenes, but also non-substituted aromatic compounds like benzene and naphthalene. The most prominent species are the toluenedegrading Desulfobacula toluolica (Rabus et al. 1993; Kuever et al. 2005a), the hexadecane- and 1-hexadecene-degrading candidatus species Desulfococcus oleovorans (Aeckersberg et al. 1991), and the species Desulfatibacillum alkenivorans (CravoLaureau et al. 2004b; Callaghan et al. 2012). Desulfosarcina cetonicum is another species known to degrade toluene (Morasch et al. 2001b; Kuever et al. 2005b), and Desulfoglaeba alkenexedens (Davidova et al. 2006) and Desulfatibacillum aliphaticivorans (Cravo-Laureau et al. 2004a) are additional validated alkane- or alkene-degrading species. Further isolates reported in pure culture but not yet taxonomically validated are listed in > Table 17.2. In addition to further toluene- and alkane/ alkene-degrading strains, they also include several deltaproteobacterial strains with newly recognized substrate degradation properties. Among these are strains oXyS1 and mXyS1, which degrade o- and m-xylene, respectively (Harms et al. 1999), the ethylbenzene-degrading strain EbS7 (Kniemeyer et al. 2003), and the naphthalene- and 2-methylnaphthalene-degrading strains NaphS2, NaphS3, and NaphS6 (Galushko et al. 1999; Musat et al. 2009). Furthermore, a sulfate-reducing Deltaproteobacterium oxidizing short gaseous alkanes like propane and butane was described, strain BuS4 (Kniemeyer et al. 2007). Whereas all strains mentioned above are affiliated with the Deltaproteobacteria, there are also some hydrocarbondegrading strains known in other phylogenetic groups of sulfate-reducing microbes. Most of these are affiliated to the Gram-positive genus Desulfotomaculum and were reported to be involved in the degradation of aromatic hydrocarbons like xylenes or biphenyl (Morasch et al. 2004; Selesi and Meckenstock 2009), but even the euryarchaeal sulfate-reducing species Archaeoglobus fulgidus was recently reported to degrade long-chain alkenes, but not alkanes (Khelifi et al. 2010).

Phototrophs Only one anoxygenic phototrophic bacterium has been reported to date to degrade toluene anaerobically, a strain of Blastochloris

Anaerobic Biodegradation of Hydrocarbons Including Methane

17

. Table 17.2 Reported sulfate-reducing hydrocarbon-degrading bacteria. Phylogenetic rank and the range of hydrocarbons degraded are given Organism

Taxon

Substrate range

Desulfobacula toluolica

Class deltaproteobacteria

Toluene

Kuever et al. (2005a)

Toluene

Kuever et al. (2005b)

PRTOL1

Toluene

Beller and Spormann (1997b)

oXyS1

Toluene, o-xylene, o-ethyltoluene

Harms et al. (1999)

mXyS1

Toluene, m-xylene, m-ethyltoluene

Harms et al. (1999)

Desulfosarcina cetonicum

Described in

EbS7

Ethylbenzene

Kniemeyer et al. (2003)

NaphS2

Naphthalene, 2-methylnaphthalene

Galushko et al. (1999)

NaphS3

Naphthalene, 2-methylnaphthalene

Galushko et al. (1999)

NaphS6

Naphthalene, 2-methylnaphthalene

Musat et al. (2009)

Desulfococcus oleovorans strain Hxd

C12–20 alkanes, alkenes

Aeckersberg et al. (1991)

Desulfatibacillum alkenivorans

Long chain alkanes, ...

Cravo-Laureau et al. (2004b)

Desulfoglaeba alkenexedens

Long chain alkanes, ...

Davidova et al. (2006)

Desulfatibacillum aliphaticivorans

Long chain alkanes, ...

Cravo-Laureau et al. (2004a)

Strain HD-3

C6–14 alkanes

Rueter et al. (1994)

Strain Pnd3

C14–17 alkanes

Aeckersberg et al. (1998)

Strain AK-01

C13–18 alkanes

So and Young (1999)

BuS4

Propane, butane

Kniemeyer et al. (2007)

Desulfotomaculum

Phylum Firmicutes

o-/m-Xylene, biphenyl

Ficker et al. (1999); Morasch et al. (2004); Selesi and Meckenstock (2009)

Archaeoglobus

Phylum Euryarchaeota

Alkenes

Khelifi et al. (2010)

sulfoviridis (Zengler et al. 1999a). This proves that metabolic conversion of anaerobic hydrocarbons into biomass is principally possible in this physiological group of bacteria, but it is not known whether phototrophs contribute very much to anaerobic hydrocarbon degradation in nature.

thermodynamic feasibility of the overall process by lowering the partial pressure of hydrogen in the shared habitat to typical values around 1 Pa (see equations below for toluene degradation). a: Toluene þ 14H2 O ! 7CO2 þ 18H2

DG0 ¼ þ504:7 KJ=mol DG0 ð1Pa H2 Þ ¼

b: 4H2 þ CO2 ! CH4 þ 2H2 O

DG0 ¼

7:9 KJ=mol

139:1 KJ=mol

Methanogenic Consortia

DG0 ð1Pa H2 Þ ¼ 25:2 KJ=mol Sum: Toluene þ 5H2 O ! 2:5CO2 þ 4:5CH4 DG 0 ¼ 121:5 KJ=mol

The first reports that anaerobic degradation of aromatic hydrocarbons occurs via disproportionation to CO2 and methane in the apparent absence of any exogenous electron acceptors almost coincided in time with those reporting hydrocarbon degradation under anaerobic respiratory conditions (Grbic-Galic and Vogel 1987; Edwards and Grbic-Galic 1994). The observed process requires the joint activities of hydrogen-degrading microorganisms capable of disposing excess reducing equivalents by formation of molecular hydrogen and methanogenic Archaea, which convert hydrogen and CO2 to methane and are responsible for the

Later studies confirmed the initial observations with aromatic hydrocarbons and showed that not only aromatic hydrocarbons but even alkanes are degraded under methanogenic conditions, both in laboratory experiments (Zengler et al. 1999b; Anderson and Lovley 2000) and in field studies (Bolliger et al. 1999; Jones et al. 2008). While none of the hydrocarbon-degrading syntrophic microorganisms are known as pure cultures or defined cocultures yet, the analysis of the prevalent organisms in enrichments or in situ populations of hydrocarbon-degrading methanogenic consortia indicates that

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Anaerobic Biodegradation of Hydrocarbons Including Methane

they may belong to several phylogenetic groups of bacteria. The expected hydrocarbon-degrading partners of the consortia are mostly affiliated with the genera Syntrophus and Smithella of the Deltaproteobacteria (Zengler et al. 1999b; Grabowski et al. 2005; Gieg et al. 2008; Jones et al. 2008), or with the genus Desulfotomaculum of the Gram-positive Firmicutes (Ficker et al. 1999). All these genera contain known strains that are involved in syntrophic degradation of fatty acids or alcohols and therefore fit well to the expected physiological role as syntrophic hydrocarbon degraders. Further phylogenetic groups of anaerobic microorganisms are often found in habitats with methanogenic oil degradation, among others a poorly characterized branch of the Chloroflexi comprising the genus Anaerolinea (Ficker et al. 1999), but it is unclear whether any of these organisms may contribute to hydrocarbon metabolism. The methanogenic partners present in the consortia seem to differ greatly dependent on the physical or chemical conditions of the habitats. It appears that hydrogenotrophic methanogens prevail mostly in habitats with high temperature, while low temperature environments may either be dominated by acetoclastic methanogens (Grabowski et al. 2005; Pham et al. 2009) or other genera of hydrogenotrophic methanogens (Grabowski et al. 2005; Shimizu et al. 2007).

Sulfidogenic Methane-Degrading Consortia The deep-sea methane seeps are the habitat of remarkable syntrophic associations of archaeal strains related to methanogens and affiliated to the order Methanosarcinales, and sulfate-reducing bacteria. These consortia are able to oxidize methane anaerobically to CO2 under sulfate-reducing conditions, despite the very low gain of free energy from the overall reaction (CH4 + SO42 + H+ ! CO2 + 2H2O + HS ; DG0 = 13.1 kJ/mol) and the high activation energy usually needed to activate methane (Boetius et al. 2000; Nauhaus et al. 2002, 2007; Knittel and Boetius 2009). The composition of the syntrophic consortia was identified by fluorescent in situ hybridization (FISH) with fluorescently labeled oligonucleotide probes targeting the 16S rRNAs of the bacterial and archaeal partners. Microscopic inspection with different probes revealed physical aggregates of the archaeal partners belonging to different phylogenetic branches within the Methanosarcinales (called ANME-1 to ANME–3 for ‘‘anaerobic methane oxidation’’) with sulfidogenic bacterial partners of the Desulfosarcina/Desulfococcus branch or the Desulfobulbus group of the Deltaproteobacteria. The different ANME partners appear to be specific for certain habitats, e.g., ANME-1 dominating the bacterial reefs in the Black Sea, or ANME-2 being the predominant archaeal partners in the associations found at Pacific gas seeps (Boetius et al. 2000; Michaelis et al. 2002). Moreover, different structures of the microbial aggregates are observed, ranging from huge archaealdominated associations interspersed with only few pockets of sulfidogenic bacteria in the Black Sea bacterial reefs (Michaelis et al. 2002) over smaller structured grains consisting of an outer shell of sulfidogenic bacteria around an inner core of tightly

packed archaeal cells prevalent at sites in the Pacific Ocean (Boetius et al. 2000) to more randomly organized consortia (Schreiber et al. 2010). No cells of any member of the ANME groups of Archaea have yet been found outside of these consortia, whereas the bacterial partners involved seem to be highly similar to free-living sulfate-reducing relatives with similar physiologies (Pernthaler et al. 2008; Schreiber et al. 2010; Basen et al. 2011). It is proposed that the ANME partners catalyze the actual methane oxidation via ‘‘reverse methanogenesis’’ (Thauer 2011), whereas the bacterial partners transfer the generated redox equivalents to sulfate. Despite significant efforts, it is still unknown how the necessary transfer of redox equivalents between the partners takes place. Recent metagenomic analyses of methane-oxidizing consortia have reinforced the importance of both partners, because the archaeal strains do not contain the genes needed for sulfate reduction (Basen et al. 2011).

Fermentative Bacteria The only truly fermentative bacterium known to be involved in anaerobic hydrocarbon degradation is the Deltaproteobacterium Pelobacter acetylenicus (Schink 1985; Ro¨sner and Schink 1995). This species represents the only known anaerobic acetylene degrader, which is quite remarkable since alkynes are virtually non-detectable in natural oil or gas deposits or any other anaerobic habitats. The bacterium can be readily enriched with acetylene as only carbon source and disproportionates acetylene to acetate and ethanol (2C2H2 + 3H2O ! H3C– COO + C2H5OH + H+; DG0 = 375 kJ/mol). It even shows relatively fast growth rates when growing on acetylene (doubling times of about 5 h), consistent with the highly exergonic overall reaction, which should allow the conversion of acetylene even at extremely low concentrations. However, the ecological significance of the anaerobic alkyne metabolism and physiological role of this bacterium are still unclear.

‘‘Intra-aerobic’’ Anaerobes Deriving O2 from Either Chlorate or Nitrite A new physiological group of anaerobic respiratory bacteria was identified in the 1990 decade, which utilize chlorate or perchlorate as electron acceptors and evolve oxygen in the course of their reduction (Malmqvist et al. 1991; Coates et al. 1999). These organisms are capable of producing oxygen for metabolic functions even in anoxic environments and are therefore referred to as ‘‘intra-aerobic’’ anaerobes. In these bacteria, perchlorate and chlorate are successively reduced by the related but distinct periplasmic molybdenum enzymes, perchlorate reductase and chlorate reductase (Coates et al. 1999; Kengen et al. 1999; Wolterink et al. 2003). The reaction product of chlorate reductase, chlorite, is immediately detoxified by chlorite dismutase, a heme-enzyme that dimutates chlorite into chloride and molecular oxygen (van Ginkel et al. 1996; Coates et al. 1999). Because this allows perchlorate- and chlorate-respiring bacteria to produce O2 even in

Anaerobic Biodegradation of Hydrocarbons Including Methane

otherwise anoxic environments, it is not very surprising that several of these ‘‘intra-aerobic’’ organisms have been reported to degrade hydrocarbons. For this purpose, they use the usual aerobic mono- or dioxygenase-dependent pathways. Strains of benzene-, toluene-, ethylbenzene-, and xylene-degrading Dechloromonas species were reported (Chakraborty et al. 2005), as well as a benzene-degrading strain of Alicycliphilus denitrificans (Weelink et al. 2008) and an n-alkane degrading strain of Pseudomonas chloritidismutans (Mehboob et al. 2009). A very different physiological group of ‘‘intra-aerobic’’ hydrocarbon-degrading bacteria was only recently identified from studies on a microbial enrichment that coupled anaerobic methane oxidation to denitrification. Anaerobic methane oxidation in this culture was apparently independent of archaeal partners and only dependent on the highly enriched main partner candidatus Methylomirabilis oxyfera, a bacterium affiliated to the novel phylum NC10 (Ettwig et al. 2010). This deeply branching bacterial phylum contained so far only bacteria known by their 16S rRNA sequences from molecular ecological studies, with Methylomirabilis oxyfera establishing the first cultured representative. Further experiments with the enrichment culture demonstrated that this organism uses an ‘‘intra-aerobic’’ strategy and evolves molecular oxygen by disproportionation of either nitrite or nitric oxide to N2 and O2 via an unknown mechanism (Ettwig et al. 2010; Wu et al. 2011). Methane is then degraded by the usual aerobic pathway, starting with a copper-dependent methane monooxygenase of the PmoA type (Ettwig et al. 2010). A similar strategy was recently proposed for the metabolism of long-chain n-alkanes by the denitrifying strain HdN1, which represents a deeply branching subgroup of the Gammaproteobacteria (Zedelius et al. 2011) (> Table 17.1). As for Methylomirabilis oxyfera, it is proposed that O2 is produced from nitrite or nitric oxide for fueling the initial activation of alkanes by monooxygenases, although no oxygen formation was detected for this strain. However, the relative amount of oxygen needed to hydroxylate long-chain alkanes is much lower than that for methane hydroxylation and therefore may be expected to be below the detection limit of the available assay (Zedelius et al. 2011). Finally, the existence of a nitrite- or NO-fueled ‘‘intra-aerobic’’ pathway of oxygen generation may also explain the puzzling observations of ‘‘anaerobic’’ degradation of many aromatic hydrocarbons by some Dechloromonas strains grown under denitrifying conditions in the absence of chlorate or perchlorate (Coates et al. 2001). The genome of one of these strains, D. aromatica strain RCB, was sequenced and contains many genes for oxygenases capable of hydroxylating aromatic compounds, but none of the known genes connected with anaerobic hydrocarbon degradation pathways (Salinero et al. 2009). Because the reactions involved in hydrocarbon metabolism in these ‘‘intra-aerobic’’ microorganisms are essentially identical to those of aerobic hydrocarbon degraders, the biochemical details are not covered in this chapter. The reader is referred to > Chap. 5, ‘‘Hydrocarbon-Oxidizing Bacteria’’ in this volume.

17

Cultivation Cultivation of anaerobic hydrocarbon-degrading bacteria or methanogenic cocultures is routinely performed in stoppered glass bottles up to the 10-L scale or in stainless steel fermentors up to scales of several 100 L. Special precautions have to be taken in cultivating sulfidogenic cultures because of the corrosive effects of the produced sulfide on steel surfaces. The media described in the following section are for the cultivation of sulfate-reducing or nitrate-reducing hydrocarbondegrading microorganisms, which are most often used in studying this topic. For use with microorganisms with other physiologies, the media can easily be modified, e.g., by replacing sulfate or nitrate with other electron acceptors or by omitting any possible electron acceptors to create optimal conditions for methanogenic consortia. The described basal media should be transparent and without organic nutrients other than the hydrocarbon substrate to be added. Just before inoculation, the media for denitrifying bacteria may be reduced by adding ascorbate as mild reductant; those for sulfate-reducing microorganisms have to be reduced by adding sodium sulfide and/or dithionite. Anaerobiosis may also be checked by adding the redox dye resazurin to the medium (1 mg/L), which is recommendable for the sulfate-reducing cultures. Components, which are heat-sensitive or lead to the formation of precipitates, are added from separate sterilized stock solutions before inoculation. Oxygen is best excluded by several cycles of evacuating and gassing with N2 in stoppered bottles (preferably with stirring or shaking after heating the solutions), but can also be avoided by alternative methods (e.g., use of anaerobic chambers). General stock solutions are prepared as follows: Trace element solution (1000x) Contains per liter

A

B

C

FeCl2 4H2O

1,500 mg 3,000 mg

None

FeSO4 7H2O

None

None

2,100 mg

MnCl2 4H2O

100 mg

100 mg

100 mg

CoCl2 6H2O

190 mg

190 mg

24 mg

Na2MoO4 2H2O

36 mg

36 mg

36 mg

ZnCl2

70 mg

70 mg

None

ZnSO4 7H2O

None

None

144 mg

CuCl2 2H2O

2 mg

4 mg

29 mg

NiCl2 6H2O

24 mg

24 mg

24 mg

H3BO4

6 mg

6 mg

30 mg None

25 % HCl

10 mL

None

NTA

None

143.3 g (adjust to pH 6.0) None

NaEDTA

None

None

5,200 mg

The FeCl2 is dissolved either in HCl (A) or in a pH-adjusted tenfold-concentrated solution of NTA (B) or NaEDTA (C), then the other components are added, and the volume is adjusted to 1 L. The solutions are autoclaved and stored at room temperature (preferably anoxically under N2 atmosphere).

615

616

17

Anaerobic Biodegradation of Hydrocarbons Including Methane

Solution (A) is left acidic to avoid precipitation of metal hydroxides and should be used with care if the media do not contain high buffer capacity. Solutions (B) and (C) contain NTA and EDTA, respectively, as potential alternative substrates and should therefore be used with an appropriate control (culture without added substrate). Trace element solutions (A) and (B) are preferably used for sulfate reducers, while trace element solution (C) is used for nitrate-reducing bacteria. Selenite/tungstate supplement (1000) Contains per liter Na2SeO3  5H2O

6 mg

Na2WO4  2H2O

8 mg

NaOH

400 mg

Autoclave and store at room temperature (preferably anoxically under N2 atmosphere). Vitamin solution (1000) Contains per liter: D(+) Biotin

20 mg

Cyanocobalamin (vitamin B12)

100 mg

p-Aminobenzoic acid

80 mg

Nicotinic acid

200 mg

Calcium pantothenate

100 mg

Thiamine-HCl  2H2O

200 mg

Pyridoxine-HCl

300 mg

Dissolve vitamins in water and filter-sterilize into a sterile screw cap bottle (0.2 mm pore size). The solution is made anoxic by degassing and inflating with N2 through a sterile filter. It is stored at 4  C in the dark. Bicarbonate Solution Dissolve 84 g NaHCO3 in deionized water to a final volume of 1 L and flush with N2 in a stoppered bottle. Autoclave and store at room temperature. Sodium Sulfide Solution Dissolve 48 g Na2S  9 H2O in 1 L of deionized water by stirring under an N2 atmosphere and autoclave in a bottle with fixed rubber or viton stopper. Sodium Dithionite Solution Dissolve 5 g of Na2S2O4 under anoxic conditions (in an anoxic glove box or in a stoppered bottle with N2 atmosphere) in 100 mL of anoxic, deionized water (e.g., from a screw cap bottle). Store in the dark for up to 2 weeks. Filter-sterilize (0.2-mm pore size) during application to the media. Sodium Ascorbate Solution Dissolve 9 g of ascorbic acid by adding 40 mL deionized water and slowly adjusting the pH to 8–9 by adding 40 mL of 1 M NaOH (preferentially on an ice bath and under N2 atmosphere). Fill up to 100 mL, filter-sterilize, and store under an N2 headspace in the dark at 4  C.

Preparation of media: Basal medium for sulfate- or nitrate-reducing bacteria, respectively, contains per liter:

NaCl

A

B

0.5 g

20.0 g

C

MgCl2  6H2O

0.5 g

3.0 g

2.0 g

CaCl2  2H2O

0.1 g

0.15

0.15 g

NH4Cl

0.3 g

0.25 g

0.54 g

KH2PO4

0.2 g

0.2 g

KCl

0.5 g

0.5 g

NaH2PO4

0.6 g

K2HPO4

5.6 g

For sulfate-reducing bacteria Na2SO4

4.0 g

4.0 g

4.0 g

For nitrate-reducing microorganisms NaNO3

0.43 g

0.43 g

0.43 g

Na2SO4

0.14 g

0.14 g

0.14 g

All media compounds are dissolved in distilled water to a final volume of 1 L. (A) and (B) are the basal formulas for bicarbonate-buffered media either for microorganisms from freshwater habitats (A) or for most marine microorganisms (B). Only occasionally, some marine isolates may need higher concentrations of MgCl2 (increase to 5.0 g/L) and CaCl2 (raise to 1.4 g/L), but these modifications also lead to increased formation of precipitates, especially at higher pH values. (C) is an alternative phosphate-buffered medium that allows growth of most freshwater isolates in the absence of added bicarbonate. The media are either supplemented with sulfate or nitrate as terminal electron acceptors to allow growth of sulfate- and nitrate-reducing microorganisms, respectively. Note that the medium for nitrate-reducing microorganisms also has to be supplemented by low sulfate concentrations for assimilatory purposes. In case of methanogenic cultures or cultures with other electron acceptors, nitrate can be omitted and sulfate reduced to the concentration needed for assimilation. Media are made anoxic in stoppered bottles by repeated cycles of evacuation and gassing with N2; then the closed bottles are autoclaved and stored at room temperature. If the medium has to be transferred to cultivation tubes in many small aliquots, it may be prepared in special flasks equipped with tubes for sterile anoxic gassing and an outlet valve, which allow sterile anoxic transfer to smaller cultivation devices (Widdel and Bak 1992). Prior to inoculating, the media need to be supplemented as follows: Per liter of basal media

(A) or (B)

(C)

NaHCO3 solution

30.0 mL

None

Trace element solution

1.0 mL

1.0 mL

Se/W solution (optional)

1.0 mL

1.0 mL

Anaerobic Biodegradation of Hydrocarbons Including Methane

Per liter of basal media

(A) or (B)

(C)

Vitamin solution

1.0 mL

1.0 mL

Reductants for sulfate-reducing bacteria Na2S solution

7.5 mL

1.0 mL

Na-dithionite solution

0.6 mL

1.0 mL

Reductants for nitrate-reducing bacteria (optional) Sodium ascorbate solution

3.0 mL

1.0 mL

Adjust the final pH of the medium to the desired value by adding small amounts of sterile 1 M H2SO4 or 1 M HCl (approximately 0.5–1 mL per liter of medium are needed to reach pH 7.0). Addition of Se and W is recommended for sulfatereducing microorganisms, which often harbor many seleno- and tungsto-enzymes, whereas these elements may be omitted for most nitrate-reducing microorganisms. The dithionite solution is filter-sterilized while applying it to the medium. Reduction of the medium with Na-ascorbate is not necessary for most nitrate-reducing microorganisms and may lead to growth of ascorbate-degrading microorganisms in enrichment cultures. The hydrocarbon substrates of interest are added separately to each culture vessel, usually at concentrations of 1 mM. More detailed procedures are described in the following section.

Cultivation Methods with Hydrocarbons Depending on the hydrocarbon substrate, one has to deal with gaseous, liquid, or solid compounds that have to be transferred into the growth media. Moreover, some hydrocarbons are toxic to microbial cells and will kill the cells if administered in too high concentrations. Gaseous Hydrocarbons. These hydrocarbons are usually purchased in steel bottles, from where they are released via gauges and a tubing system. They are sterilized by passing the gas through sterile cotton or membrane filters, while injecting it in the headspace of the culture bottles via hypodermic needles. The needles should be preflushed with the hydrocarbon gas to remove residual oxygen. The added amount is proportional to the added volume (molar volume of gasses equals to 22.4 mL per mmol at standard conditions). A more precise way of adding known amounts of gaseous substrates is to fill an evacuated sterilized bottle with the gas and to transfer defined amounts from there to the culture vessels by gas-tight sterile syringes. The application of high pressure of the added hydrocarbon was only reported to be beneficial for anaerobic oxidation of methane (Nauhaus et al. 2002), whereas no differences are known for higher hydrocarbons. Liquid Hydrocarbons. These compounds can be sterilized by filtration through solvent-resistant cellulose filters (pore size 0.2 mm) or by autoclaving in tightly closed bottles (check for losses for volatile compounds). Screw-cap bottles with Teflon-coated seals (> Fig. 17.5) are useful for autoclaving and storage. A special glass flask has been reported to be useful for sterilization, storage, and distribution of crude oil without loss

17

of volatile compounds (Rabus and Widdel 1996) (> Fig. 17.5). Hydrocarbons may easily be transferred from the stocks to the media using anoxic (N2-gassed) syringes (> Fig. 17.5). In special cases, care should be taken to avoid potential contamination of the hydrocarbon substrates by compounds from stoppers or plastic materials (e.g., use glass syringes, avoid rubber-sealed plungers, avoid contact of the hydrocarbon phase with the stoppers of the culture vessels; see > Fig. 17.5). These special precautions can usually be alleviated when the degradation of a substrate is established and potential effects originating from contaminations have been excluded. Most liquid hydrocarbons are poorly soluble in water and float at the surface of the culture. Therefore, it is often beneficial for growth to incubate the vessels horizontally to get a large surface area (> Fig. 17.5) and to shake or stir the media to increase the contact of the cells to the hydrocarbon droplets. However, too fast shaking may also lead to cell damage and death of the culture. Therefore, the optimal cultivation methods must be developed individually for each strain and substrate. Another problem often encountered is the toxicity of hydrocarbons. For example, toluene is soluble in water up to 5 mM but is already toxic for T. aromatica at concentrations of 1– 2 mM. This can be counteracted by adding only small amounts of the compounds to keep the concentration below the toxicity level and refeed the culture when the initial amount has been consumed, which is very labor-intensive and needs constant inspection of the cultures. A much easier alternative approach is to establish a two-phase system with an inert hydrophobic phase that serves as carrier phase and reservoir for the hydrocarbon substrate (> Fig. 17.5). Because the hydrocarbon is also equilibrating with the aqueous phase, it will be available in small, nontoxic concentrations for the degrading microorganisms and re-equilibrate from the hydrophobic reservoir in parallel to the degradation rate. Compounds usually used as carrier phases are paraffin oil, pristane, or 2,2,4,4,6,8,8-heptamethylnonane. Paraffin oil is relatively low-prized compared to the other alternatives, but cannot be used for enrichments or for the cultivation of alkane-degrading microorganisms, because it is composed of long-chain alkanes itself. An alternative method to establish a two-phase system with a solid XAD7 resin as hydrophobic phase was reported for degradation of aromatic compounds by sulfate-reducing bacteria (Morasch et al. 2001a). Solid Hydrocarbons. Several hydrocarbons (e.g., naphthalene) are solid at room temperature. Media containing these compounds as carbon source may be prepared by adding a piece of the solid compound to each medium bottle. A possible alternative is to prepare a stock solution of the solid hydrocarbon in an inert liquid carrier phase (e.g., paraffin or tert-butanol) and distribute it like liquid hydrocarbons.

Biochemistry of Microbial Hydrocarbon Degradation The biochemistry behind anaerobic hydrocarbon degradation has been studied in detail for the BTEX compounds

617

618

17

Anaerobic Biodegradation of Hydrocarbons Including Methane

a

b Sterile N2 Screw cap with Teflon-coated sealing disk

Syringe

Screw stop cock with Teflon plunger

Liquid hydrocarbon

c

Screw cap with Teflon-coated sealing disk

Liquid hydrocarbon

Liquid hydrocarbon Medium

Medium

. Fig. 17.5 Supplementation and feeding of anoxic culture media. Storage of (sterile) hydrocarbons and their addition to anoxic culture medium. (a) Use of screw bottle with stopper or sealing disk. (b) Advanced method using a special bottle with additional access for supplementation and feeding. (c) Contact of hydrocarbon or hydrophobic carrier phase with the stopper can be avoided during injection and incubation. Moreover, horizontal incubation of the vessels and a thereby large surface area can increase substrate accessibility

(benzene, toluene, ethylbenzene, and the three xylene isomers), the bi-aromatic compounds naphthalene and methylnaphthalene, and short- and long-chain alkanes. Degradation pathways for these compounds have been proposed and are supported by the identification of key intermediates, stable isotope analysis, biochemical studies, as well as proteomic and genomic data. Six different general catalytic principles for initial reactions with different hydrocarbons are firmly recognized. (1) The addition of fumarate to methyl or methylene groups of the hydrocarbons leads to the formation of succinate adducts as primary intermediates. This reaction is catalyzed by a novel group of glycyl radical enzymes. (2) An oxygen-independent hydroxylation of alkyl chains to secondary or tertiary alcohols is catalyzed by molybdenum-cofactor containing periplasmic dehydrogenases. (3) Activation of hydrocarbons by direct carboxylation is proposed to be catalyzed by unique carboxylases of a novel

enzyme family. (4) Hydration of acetylene is catalyzed by a special tungsten-cofactor containing enzyme, acetylene hydratase. (5) Reversible hydration of C = C double bonds of terpenoid hydrocarbons is catalyzed by a novel type of hydratase. (6) The anaerobic oxidation of methane apparently follows a unique pathway of ‘‘reverse methanogenesis’’ that is physiologically linked to the metabolic activities of at least two prokaryotic partners in syntrophic associations. In the following section, the biochemical background of these processes is described.

Fumarate Addition Reactions Toluene. The anaerobic toluene metabolic pathway represents a model for the widely distributed type of initiation reaction via

Anaerobic Biodegradation of Hydrocarbons Including Methane

BssD (act.)

CH3

tdiR

COO-

BssABC

Fumarate Toluene

tdiS

QH2

COO-

bssD bssC

Q COO-

bssA

COO-

bssB bssE

COOSuccinate

(R)-Benzylsuccinate

bss operon

Sdh

COO-

17

bssF O

SCoA

COSCoA

BbsEF

COSCoA

COOCOOSuccinyl-CoA

BenzoylCoA

BbsAB CoASH

bssH ETFox bbsG

BbsG

(R)-Benzylsuccinyl-CoA

COSCoA

bssG

bbsH

ETFred

bbsJ

COSCoA

COO-

COO-

Benzoylsuccinyl-CoA

COSCoA

(E)-Phenylitaconyl-CoA

BbsCD Bb

COO-

NAD+

bbsF bbsE bbsD bbsC

HO NADH

bbs operon

O

H2O BbsH

bbsB bbsA

. Fig. 17.6 Anaerobic degradation of toluene by initial addition of fumarate. The initial reaction leads to the formation of (R)-benzylsuccinate, catalyzed by benzylsuccinate synthase (BSS). BSS is a glycyl radical enzyme that needs to be activated by a specific activating enzyme (BssD). The genes coding for the subunits of BSS (bssABC) and activating enzyme (bssD) form an apparent operon together with several genes of unknown function (bssE-H). (R)-benzylsuccinyl-CoA is further metabolized via a beta-oxidation pathway. The genes coding for the enzymes involved are located in a second apparent operon (bbsA-G). The enzymes catalyzing the different steps of the pathway are indicated by the respective gene products. Both operons are regulated coordinately by the specific two-component regulation system, TdiSR

fumarate addition (Biegert et al. 1996; Beller and Spormann 1998; Heider et al. 1998; Spormann and Widdel 2000; Widdel and Rabus 2001; Chakraborty and Coates 2004), which also occurs with many further hydrocarbons, and was intensively studied in the last decade. The capability for anaerobic toluene metabolism is widespread, which may be explained by the general abundance of toluene and its relatively high solubility in water. All anaerobic toluene-degrading bacteria known to date, which represent all known physiological groups of anaerobic hydrocarbon degraders, initialize degradation of toluene via the addition of a fumarate cosubstrate to form benzylsuccinate (Rabus and Heider 1998; Zengler et al. 1999a; Beller and Edwards 2000; Kane et al. 2002) (> Fig. 17.6). The detailed biochemistry of anaerobic toluene metabolism has been studied in denitrifying Thauera and Azoarcus (Aromatoleum)

species. The initial step, stereospecific activation of toluene to (R)-benzylsuccinate, is catalyzed by the extremely oxygen-sensitive glycyl radical enzyme benzylsuccinate synthase (BSS), which was purified from Thauera aromatica and Azoarcus strain T (Leuthner et al. 1998; Beller and Spormann 1999). Other known glycyl radical enzymes are pyruvate formate lyase (PFL), anaerobic ribonucleotide reductase (Nrd), coenzyme B12-independent glycerol dehydratase (GD), and 4-hydroxyphenylacetate decarboxylase (HPD) and some related anaerobic decarboxylases (Selmer et al. 2005). They are generally homodimeric enzymes with large subunits of 80–100 kDa, which need to be activated by separate S-adenosyl-methionine (SAM)-dependent activating enzymes that belong to the large family of ‘‘SAM-radical’’ enzymes. It has been proven that in its activated state, BSS carries a stable glycyl radical at the conserved

619

620

17

Anaerobic Biodegradation of Hydrocarbons Including Methane

glycyl residue of the catalytic center, as indicated by EPR spectroscopy and by identification of a characteristic cleavage product of the glycyl radical site of the polypeptide chain after inactivation by oxygen (Leuthner et al. 1998; Krieger et al. 2001; Verfu¨rth et al. 2004). A second important part of the catalytic center of BSS (as of other glycyl radical enzymes) is a conserved cysteine residue that is predicted to be localized close enough to the glycyl radical site in the tertiary structure to be activated to a transient thiyl radical species (Selmer et al. 2005). A catalytic mechanism for benzylsuccinate synthesis has been proposed based on these findings, which is initiated by abstracting a hydrogen atom from the methyl group of toluene, leading to the formation of a substrate-related benzyl radical intermediate (Heider et al. 1998). The benzyl radical formed immediately attacks the double bond of a fumarate cosubstrate supplied in the active center, leading to a product-related radical intermediate that regains the hydrogen atom from the enzyme, forming benzylsuccinate and restoring the radical form of the enzyme. The proposed mechanism is in agreement with ab initio calculations (Himo 2002, 2005) and with studies with deuterium-labeled toluene that show that the initially abstracted hydrogen atom of toluene is conserved in benzylsuccinate (Beller and Spormann 1997a, b, 1998). In addition to the large glycyl-radical containing subunit of 98 kDa, BSS contains two unique small subunits (8.5 and 6.5 kDa, respectively), which apparently contain [Fe4S4]-clusters of very low midpoint potential (Li et al. 2009; Heider and Boll 2010) and seem to be essential for catalysis (unpublished results). The only other known glycyl radical enzyme with additional [Fe4S4]-clusters is HPD, which also carries these clusters in an additional small subunit unique to this subfamily of enzymes (Selmer et al. 2005). Like the other glycyl radical enzymes, BSS is synthesized as inactive precursor protein and needs a specific SAM-dependent activating protein, which is encoded in the same operon (Leuthner et al. 1998; Hermuth et al. 2002). In addition to the genes for the three subunits of the enzyme (bssABC) and the gene for the activating enzyme (bssD), the known operons for BSS and other enzymes of fumarateadding glycyl radical enzymes contain up to four more conserved genes for proteins of unknown function (bssEFGH), which may be involved in the assembly or the activation of the enzymes (Hermuth et al. 2002; Kube et al. 2004; Rabus et al. 2005) (> Fig. 17.6). Benzylsuccinate is further metabolized by activation to benzylsuccinyl-CoA and further degradation via a b-oxidation reaction sequence, leading to the formation of benzoyl-CoA and succinyl-CoA (Leutwein and Heider 1999, 2002) (> Fig. 17.6). All enzymes involved in this pathway are encoded in a second operon (bbsA-H) (Leuthner and Heider 2000). Succinyl-CoA is used as CoA donor for benzylsuccinate activation by a CoA transferase (BbsEF) (Leutwein and Heider 2001), and the produced succinate is oxidized by succinate dehydrogenase to restore the fumarate cosubstrate. Both the bss and the bbs operons are coordinately induced in denitrifying bacteria via a two-component regulatory system (TdiSR) only in the presence of toluene and the absence of oxygen

(Leuthner and Heider 1998). The product of the b-oxidation pathway, benzoyl-CoA, is the central intermediate of anaerobic metabolism of aromatic compounds and is further degraded via reduction of the aromatic ring by benzoyl-CoA reductase and further hydrolytic and b-oxidation-like reactions (described in detail in Harwood et al. (1998); Boll et al. (2002); Carmona et al. (2009); Heider and Boll (2010)). Benzylsuccinate synthases from toluene-degrading bacteria exhibit quite broad substrate spectra for activation of other aromatic compounds, which vary between the enzymes of different strains (Verfu¨rth et al. 2004). Interestingly, the enzymes catalyze fumarate addition to a variety of aromatic compounds that are not degraded by the respective strains, and in some cases the derived succinate adducts have been observed as dead-end products in the medium (Beller and Spormann 1997a). Therefore, the degradation capacity of a bacterial strain via fumarate addition seems to be determined by the substrate specificities of both the benzylsuccinate synthases and the enzymes of the corresponding b-oxidation pathway(s) present in the respective strain. In natural environments, the broad substrate spectrum of benzylsuccinate synthases could lead to cross-feeding of succinate adducts to other members of the bacterial community, but may also account for the production of dead-end succinate adducts from many petroleum constituents. These metabolites are indeed often detected together with benzylsuccinate in contaminated aquifers, but it is not clear whether or how fast they are degraded further to CO2 (Beller and Spormann 1997a; Elshahed et al. 2001; Rios-Hernandez et al. 2003; Griebler et al. 2004; Parisi et al. 2009). Clear evidence for complete degradation of the initially formed succinate adducts is only available for a few hydrocarbons, which are listed below. Xylenes. All three xylene isomers (o-/m-/p-) as well as the cresol isomers and o-toluidine were shown to be activated by ‘‘normal’’ (toluene-induced) benzylsuccinate synthases via fumarate addition (Beller and Spormann 1997a; Krieger et al. 1999; Verfu¨rth et al. 2004; Morasch and Meckenstock 2005). Of the xylenes, m-xylene is most readily degraded, and several pure cultures of denitrifiers and sulfate reducers have been isolated that grow almost equally well with toluene or m-xylene (Spormann and Widdel 2000; Widdel and Rabus 2001). Pure cultures degrading o-xylene are rare, and so far only two sulfate-reducing strains have been described to completely utilize o-xylene to CO2 (Harms et al. 1999; Morasch et al. 2004). The complete degradation of p-xylene to CO2 has recently been reported with enrichment cultures dominated by members of the Rhodocyclaceae, but only distantly related to the genera Thauera and Aromatoleum (Rotaru et al. 2010). In all cases, succinate adducts of the respective hydrocarbons are detectable in the culture supernatants (Krieger et al. 1999; Morasch et al. 2004; Rotaru et al. 2010), suggesting the involvement of the same or very similar BSS isoenzymes as for toluene conversion. 2-Methylnaphthalene. Degradation of the polyaromatic hydrocarbon 2-methylnaphthalene (> Fig. 17.1) by a sulfatereducing enrichment culture was shown to be initiated via fumarate addition to the methyl side chain of the hydrocarbon

Anaerobic Biodegradation of Hydrocarbons Including Methane

(Annweiler et al. 2000; Selesi et al. 2010). Moreover, the genes coding for the subunits of the naphthyl-2-methyl-succinate synthase (nmsABC) and its associated activating enzyme (nmsD) were identified. The enzymes show high similarity with the respective subunits of BSS and BSS-activating enzymes while forming a separate branch of the fumarate-adding glycyl radical enzyme subfamily. Furthermore, a second gene cluster coding for the enzymes of a b-oxidation pathway (bnsA-H) analogous to those identified for toluene degradation was described. Overall, the enzymes of this pathway catalyze the degradation of 2-methylnaphthalene to 2-naphthoyl-CoA, which is subsequently reductively dearomatized by a 2-naphthyl-CoA reductase (Ncr), in analogy to benzoyl-CoA (Harwood et al. 1998; Annweiler et al. 2000; Selesi et al. 2010). Alkanes and Cycloalkanes. n-Alkanes are degraded by pure cultures of many denitrifying or sulfate-reducing bacteria (Aeckersberg et al. 1991; Rueter et al. 1994; So and Young 1999; Ehrenreich et al. 2000; Cravo-Laureau et al. 2005) (> Tables 17.1, > 17.2). The substrate spectra vary in different strains, and the range of n-alkanes utilized extends from shortchain alkanes (propane, butane) to medium- and long-chain alkanes (C6–C20). In most strains, alkane degradation was found to be initiated by addition of fumarate to the subterminal methylene group of the respective alkanes, leading to the formation of branched (1-methylalkyl)succinates (Rabus et al. 1999, 2001; Kropp et al. 2000; Cravo-Laureau et al. 2005). Moreover, the presence of an alkane-induced glycyl radical enzyme was detected by EPR spectroscopy (Rabus et al. 2001). Based on

proteomic studies with the n-hexane-degrading denitrifying strain HxN1 and the identification of genes coding for induced proteins in hexane-grown cells, an apparent operon was revealed to code for the subunits of the expected (1-methylalkyl)succinate synthase (masABC), its associated activating enzyme (masG) and further paralogs of conserved proteins known from the operons for other fumarate-adding enzymes (Grundmann et al. 2008). The availability of the first genome sequence of a sulfate-reducing alkane degrader using fumarate addition, Desulfatibacillum alkenivorans, revealed the presence of even two separate operons for alkane-activating fumarate-adding glycyl radical enzymes (ass-1 and ass-2) (Callaghan et al. 2010, 2011). The sequences of these enzymes show again high similarity to the other known fumarate-adding glycyl radical enzymes, but establish the alkane-specific enzymes as a third separate subfamily (Callaghan et al. 2010). The branched intermediates generated from subterminal fumarate addition to the alkanes cannot directly undergo b-oxidation, because the methyl group blocks one of the carbon atoms to be oxidized. Consequently, the further degradation pathway involves a carbon skeleton rearrangement reaction at the level of CoA thioesters to move the obstacle by one carbon atom. The reaction appears to be similar to the reaction catalyzed by methylmalonyl-CoA mutase (> Fig. 17.7) and is followed by a decarboxylation at the beta-carbon atom. The resulting 4-methyl-branched fatty-acyl-CoA intermediate is feasible for b-oxidation to acetyl-CoA units and one propionyl-CoA. Investigation of intermediates present in culture supernatants

2 [H] -OOC

COO-

-OOC

Fumarate

COO-

-OOC

COO-

Succinate

CO

Succinyl-CoA

SCoA

CO

Alkane (C2n)

Methyl-alkylsuccinate

Propionyl-CoA

Methylmalonyl-CoA

R

R COO-

SCoA

[CO2]

R -OOC

CO

SCoA

[CoA] R

17

-OOC

CO

Methyl-alkylSuccinyl-CoA

SCoA

R

-OOC

[β -Ox.] CO

SCoA

(2-Methylalkyl)malonyl-CoA

CO

SCoA

4-Methylalkyl-CoA n Acetyl-CoA + (4n + 2) [H]

. Fig. 17.7 Anaerobic degradation of n-alkanes by initial addition of fumarate. The initial reaction forms a branched 1-methyl-1-alkyl-succinate adduct. The further metabolism is proposed to proceed via formation of the CoA thioester and its rearrangement to (2-methylalkyl)malonyl-CoA, which is decarboxylated to a 4-methylalkyl-CoA derivative. The latter can be beta-oxidized to yield several acetyl-CoA units and one propionyl-CoA. It has been proposed that propionyl-CoA is used to regenerate the fumarate cosubstrate. We suggest that activation of the succinate adduct and decarboxylation of (2-methylalkyl)-malonyl-CoA may occur in coupled reactions with corresponding steps of fumarate regeneration. These suggested CoA- and CO2-transfer reactions would allow an energy-efficient pathway for activation of the succinate adduct and fumarate regeneration. The metabolic fate of an incoming fumarate moiety over one cycle is indicated in red

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of alkane-degrading cultures indeed confirmed the presence of most of the proposed intermediates of this pathway (Wilkes et al. 2002). The propionyl-CoA unit produced from the methylbranched moiety of the 4-methyl-acyl-CoA intermediate is regenerated to the initial cosubstrate fumarate. This involves carboxylation to methylmalonyl-CoA and rearrangement to succinyl-CoA, followed by hydrolysis to succinate and oxidation to fumarate (Wilkes et al. 2002). It is plausible to assume that analogous steps of (1-methylalkyl)succinate degradation and fumarate regeneration occur as coupled reactions, especially CoA transfer from succinyl-CoA to (1-methylalkyl)succinate and transfer of the carboxy group from (2-methylalkyl)malonylCoA to propionyl-CoA, as shown in > Fig. 17.7. A slightly modified version of fumarate addition to alkanes was observed in the marine sulfate-reducing bacterial strain BuS5, the first pure culture known to degrade propane or butane anaerobically. During growth on propane, both iso- and n-propylsuccinate were detected as intermediates, indicating that activation of propane occurs via addition of fumarate to the subterminal methylene group as well as to the terminal methyl group (Kniemeyer et al. 2007). Moreover, previous reports on anaerobic cyclohexane degradation (Rios-Hernandez et al. 2003) have recently been verified in a nitrate-reducing bacterial consortium dominated by a Geobacter-like strain in an apparent syntrophic coupling to anaerobic ammonium oxidation (‘‘Anammox’’). The authors identified cyclohexyl-succinate in the culture supernatant as well as cyclohexyl-substituted fatty acids in the cells. This indicates that cyclohexane is activated by fumarate addition to one of the methylene carbons in the alicyclic ring and further degraded by an analogous pathway as proposed for n-alkane degradation (Musat et al. 2010). Ethylbenzene. Although ethylbenzene is known to be degraded via an oxygen-independent hydroxylation reaction in denitrifying bacteria (see below), the recently described sulfate-reducing strain EbS7 initiates ethylbenzene metabolism by fumarate addition, forming (1-phenylethyl)succinate. In analogy to the metabolism of alkanes, this branched intermediate is proposed to be further degraded by carbon-skeleton rearrangement and decarboxylation (Kniemeyer et al. 2003).

Oxygen-Independent Hydroxylation Ethylbenzene. Ethylbenzene has highly similar chemical properties as toluene and is degraded via an analogous fumarate addition at least in one known sulfate-reducing bacterium (see above). However, another completely different type of reaction was shown to initialize anaerobic ethylbenzene degradation in denitrifying bacteria, namely, an oxygen-independent hydroxylation at carbon-1 (C1) of the side chain to yield (S)-1-phenylethanol (Ball et al. 1996; Rabus and Heider 1998; Kniemeyer and Heider 2001a) (> Fig. 17.8). The reaction is strictly stereospecific and represents the first example for an anaerobic hydroxylation of a nonactivated hydrocarbon with water as donor for the hydroxyl group. The reaction is catalyzed by the periplasmatic

molybdenum/iron-sulfur/heme-enzyme ethylbenzene dehydrogenase (EBDH), which consists of three subunits and belongs to the DMSO subfamily of molybdenum enzymes (Johnson et al. 2001; Kniemeyer and Heider 2001a). The crystal structure of the reduced form of EBDH from Aromatoleum aromaticum strain EbN1 was solved, showing the structure of the active site at the molybdenum cofactor in the large subunit and the extent of an electron transfer chain through the entire enzyme, which consists of five iron-sulfur clusters in the a- and b-subunits and a heme b cofactor in the g-subunit (Kloer et al. 2006). A relatively wide channel leading to the active site may explain the very broad substrate spectrum observed for EBDH (Szaleniec et al. 2007). The physiological electron acceptor for EBDH is most probably cytochrome c, based on its periplasmic location and the apparent requirement for artificial electron acceptors of rather high potentials for enzyme assays (p-benzoquinone or ferrocenium tetrafluoroborate) (Kniemeyer and Heider 2001a). A mathematical model of a plausible reaction mechanism for the biochemically unusual oxygen-independent hydroxylation of ethylbenzene at the molybdenum cofactor was developed, which is corroborated by experimental data on the predicted isotope effects of the reaction (Szaleniec et al. 2010). For the further part of the metabolic pathway, (S)-1phenylethanol needs to cross the cytoplasmic membrane (probably by passive diffusion) and is subsequently further oxidized to acetophenone by a stereospecific NAD-dependent alcohol dehydrogenase (Kniemeyer and Heider 2001b; Ho¨ffken et al. 2006). The genes coding for the subunits of EBDH (ebdABC) and (S)-1-phenylethanol dehydrogenase (ped) are present in a common operon, which also contains an additional gene coding for a chaperone-like protein (ebdD) required for insertion of the molybdenum cofactor into EBDH and its secretion into the periplasm via the ‘‘twin-arginine transport’’ (tat) system, as inferred from other proteins of the DMSO reductase family (McDevitt et al. 2002; Rabus et al. 2002). The intermediate acetophenone is carboxylated by a novel type of ATP-dependent carboxylase, acetophenone carboxylase (APC), a complex enzyme consisting of five subunits (ApcA-E) to yield benzoylacetate (> Fig. 17.8) (Jobst et al. 2010), which is activated to the CoA thioester by a specific benzoylacetate-CoA ligase (BAL). The generated thioester is then finally cleaved to acetyl-CoA and benzoyl-CoA by a thiolase, and benzoyl-CoA is further degraded by the known central pathway of anaerobic aromatic metabolism. The genes coding for the subunits of acetophenone carboxylase and benzoylacetate-CoA ligase form another common operon (Rabus et al. 2002). The two operons involved in anaerobic ethylbenzene metabolism are regulated by two specific two-component regulatory systems in a sequential mode: whereas the ebd-ped-operon is induced under anaerobic conditions in the presence of ethylbenzene, the apc-bal-operon is induced independently of oxygen only by the presence of acetophenone (Ku¨hner et al. 2005) (> Fig. 17.8). This is consistent with the observation that ethylbenzene is only degraded in the absence of oxygen, while acetophenone degradation is independent of the redox status (Rabus and

Anaerobic Biodegradation of Hydrocarbons Including Methane

HO

2 [H]

17

H2O

EBDH Ethylbenzene

(S)-1-Phenylethanol

HO NAD+

NADH H+

Periplasm

O

2 ADP CO2 2 ATP 2 Pi

PDH (S)-1-Phenylethanol

O

COO-

Cytoplasm

APC Benzoylacetate

Acetophenone

ATP HSCoA CO~

SCoA

CH3 Acetyl-CoA CO~ SCoA

AMP PPi O

BCL

CO~ SCoA

HSCoA Benzoylacetyl-CoA

Benzoyl-CoA

. Fig. 17.8 Anaerobic degradation of ethylbenzene by initial hydroxylation in nitrate-reducing bacteria. Ethylbenzene is initially stereospecifically hydroxylated to (S)-1-phenylethanol by the periplasmatic molybdenum/iron-sulfur/heme-enzyme ethylbenzene dehydrogenase (EBDH). (S)-1-Phenylethanol is further metabolized to benzoylacetate by cytosolic enzymes, (S)-1-phenylethanol dehydrogenase (PDH) and acetophenone carboxylase (APC). Benzoylacetate is activated by a specific CoA ligase (BCL) to benzoylacetyl-CoA, which is thiolytically cleaved to acetyl-CoA and benzoyl-CoA. The genes coding for the first four enzymes of the pathway are located in two apparent operons, both of which are under the control of specific two-component regulation systems (AdiSR and EdiSR, respectively). ebdABC: genes coding for the subunits of EBDH. ebdD: gene encoding a private chaperone for EbDH assembly and export. apcABCDE: genes encoding the subunits of acetophenone carboxylase. bcl: gene encoding benzoylacetate-CoA ligase

Widdel 1995). The thiolase catalyzing the last step is not encoded in any of the operons and probably represents the general enzyme also involved in thiolytic cleavage reactions during the further metabolism of benzoyl-CoA (Harwood et al. 1998; Boll et al. 2002; Heider and Boll 2010). Propylbenzene. A bacterial isolate closely related to A. aromaticum, strain PbN1, is able to degrade both ethylbenzene and n-propylbenzene under denitrifying conditions. Because the EBDH orthologs from both strains were shown to hydroxylate both hydrocarbons with sufficient activities (Rabus and Widdel 1995; Kniemeyer and Heider 2001a), the catabolic pathway of n-propylbenzene was proposed to be analogous to that of ethylbenzene, leading to the formation of propionyl-CoA and benzoyl-CoA. Cholesterol. Although cholesterol is not a true hydrocarbon, it is mentioned here, because a paralogous enzyme highly similar to EBDH was recently identified to hydroxylate the isoprenoid alkyl side chain of the apolar ‘‘hydrocarbon-like’’ side of

cholesterol and several related steroids to yield a tertiary alcohol intermediate (Chiang et al. 2007, 2008). This oxygenindependent hydroxylation contributes to a novel pathway of anaerobic degradation of cholesterol in the Betaproteobacterium Sterolibacterium denitrificans, which is closely related to the genera Thauera and Aromatoleum. Putative Alternative Alkane Metabolic Pathway. The sulfatereducing n-alkane-degrading candidatus species Desulfococcus oleovorans (Aeckersberg et al. 1991) and a nitrate-reducing alkane-degrading consortium (Callaghan et al. 2009) are known to utilize a different pathway for alkane degradation, compared to most other alkane-degrading strains, which initiate the pathway by fumarate addition. This is evident from the finding that cells grown on even-chain alkanes contain predominantly odd-chain fatty acids (and vice versa), whereas none of the fumarate-adding strains shows this feature (Aeckersberg et al. 1998; So et al. 2003; Callaghan et al. 2006). Based on labeling studies of cellular fatty acids with 13CO2, it has been

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H2O OH

CO2

2[H] CoASH [ATP]

COO-

O

2 [H] CO2

O H2O

COSCoA

4 [H] O

COSCoA

COO-

CoASH [ATP]

CoASH

Ac-SCoA+ COSCoA

. Fig. 17.9 Possible pathways of n-alkane degradation in the sulfate-reducing bacterium Desulfococcus oleovorans; left: ‘‘old’’ pathway initiated by carboxylation, as suggested by So and Young (2002); right: alternative pathway initiated by oxygen-independent hydroxylation. Note that both putative pathways converge at the same intermediates. The fate of the CO2 utilized for carboxylation is highlighted by coloring

proposed that D. oleovorans may initiate n-alkane metabolism by carboxylating the alkanes at carbon-3 (C3), leading to 1-ethyl-branched fatty acids that may be degraded further by b-oxidation (> Fig. 17.9) (So et al. 2003). However, another possible scenario consistent with the available labeling patterns may be an initial subterminal hydroxylation of the alkanes by an EBDH-like enzyme, followed by the oxidation to a ketone and its subsequent carboxylation at C3 (> Fig. 17.9). It remains to be shown which one of the two possible paths is really followed in D. oleovorans, but it may be of interest that an operon coding for an EBDH-like enzyme is indeed present in the genome of D. oleovorans (Acc. No. YP001528081-YP001528084), which is missing in the genome sequences of other anaerobic alkane-degrading bacteria (our unpublished observations).

Carboxylation Benzene. Many enrichment cultures metabolizing benzene under anaerobic conditions have been reported with various terminal electron acceptors (Edwards and Grbic-Galic 1992; Lovley et al. 1994, 1995; Burland and Edwards 1999; Lovley 2000; Kunapuli et al. 2007; Musat and Widdel 2008; Abu Laban et al. 2009), but only few pure cultures are available to date (Coates et al. 2001; Kasai et al. 2007). Benzoate was detected as metabolite of benzene-grown cultures (Caldwell and Suflita 2000; Phelps et al. 2001; Chakraborty and Coates 2005). Yet benzoate or benzoyl-CoA is the central intermediate of anaerobic degradation of aromatic compounds and therefore also to be expected as intermediate in other proposed pathways. A recent proteomic study with an iron-reducing enrichment culture demonstrated benzene-induced expression of several

proteins that show sequence similarity with the a-subunit of phenylphosphate carboxylase described for anaerobic phenol degradation in denitrifying bacteria of the Thauera/Azoarcus cluster of Betaproteobacteria (Schu¨hle and Fuchs 2004; Abu Laban et al. 2010). Furthermore, additional carboxylase-related genes and a gene encoding a putative benzoate-CoA ligase were specifically expressed (> Fig. 17.10). Therefore, a direct carboxylation of benzene to benzoate was suggested as initial step. Naphthalene, Phenanthrene, and Biphenyl. Initial carboxylation of naphthalene and phenanthrene to 2-naphthoate and phenanthrene carboxylic acid, respectively, was first described by Zhang and Young (1997). They reported incorporation of 13 CO2 in both substrates by sulfate-reducing consortia, which completely mineralized the polycyclic hydrocarbons. Studies with pure cultures of the naphthalene-degrading deltaproteobacterial strain NaphS2 confirmed that 2-naphthoate is an intermediate of naphthalene degradation (Galushko et al. 1999; Musat et al. 2009). A cluster of highly induced genes was identified, some of which show similarity to genes coding for subunits of a known carboxylase involved in anaerobic phenol degradation (Schu¨hle and Fuchs 2004; Bergmann et al. 2011; Meckenstock and Mouttaki 2011) (> Fig. 17.10). Also, in anaerobic degradation of biphenyl by a sulfate-reducing enrichment culture, biphenylcarboxylic acid was detected as major metabolite (Selesi and Meckenstock 2009). Therefore, a pathway initiated by direct carboxylation was suggested for both polycyclic hydrocarbons. Alkanes. A mechanism for alkane degradation without fumarate addition reactions is known to operate in the sulfate-reducing bacterium Desulfococcus oleovorans and a nitrate-reducing consortium (see above) (So et al. 2003; Callaghan et al. 2006, 2009). Based on labeling experiments with 13CO2, it has been proposed that the alkane is carboxylated at C3 (So et al. 2003), but it is not sure whether this carboxylation represents the initial reaction. Two possible alternative scenarios of alkane degradation initiated either by carboxylation or by hydroxylation are shown in > Fig. 17.9.

Acetylene Hydration Acetylene hydratase of Pelobacter acetylenicus was until recently the only enzyme catalyzing water addition to an unsaturated hydrocarbon, namely acetylene. The initially formed C2-enol intermediate is immediately tautomerised to acetaldehyde. The enzyme contains a tungsten-bis-molypdopterin-guanine dinucleotide cofactor and a [Fe4S4] cluster and belongs to the DMSOreductase family of molybdenum enzymes (Ro¨sner and Schink 1995; Meckenstock et al. 1999a). It is the only member of this family catalyzing a non-redox reaction and is only active in the fully reduced W(IV) state. The recently solved structure (Seiffert et al. 2007) combined with biochemical analysis of the wild type and mutant versions of the enzyme (tenBrink et al. 2011) allows detailed studies of the mechanism of acetylene hydration, which are further backed up with mathematical modeling of potential reaction mechanisms (Liao et al. 2010; Liu et al. 2011).

Anaerobic Biodegradation of Hydrocarbons Including Methane

a ATP

OHCO2

AMP Pi O

PpsABC O

OH

P

O-

COOPi

Phenylphosphate

OH

OH

p -Hydroxybenzoate

p -Hydroxybenzoyl-CoA

COO-

b CO2 AbcAD (?) Benzene

c

CO~ SCoA

PpcABCD

OH

Phenol

AMP PPi

ATP HSCoA

17

AMP PPi

ATP HSCoA Blz

Benzoate

CO2 NacAB (?) Naphthalene

CO~ SCoA

Benzoyl-CoA

COOH ATP HSCoA

CO~ SCoA AMP PPi

. Fig. 17.10 Anaerobic degradation of phenol, benzene, and naphthalene by initial carboxylation. (a) Anaerobic phenol degradation by initial conversion to phenylphosphate by phenylphosphate synthase (PpsABC), followed by carboxylation to 4-hydroxybenzoate by phenylphosphate carboxylase (PpcABCD). The genes coding for the subunits of the respective enzyme are shown below the reactions. Two additional genes coding for putative (de)carboxylases of unknown function similar to UbiD and UbiX, two isofunctional enzymes involved in ubiquinone biosynthesis, are also encoded in the operon. (b) Proposed pathway of benzene degradation via carboxylation to benzoate and activation to benzoyl-CoA. The genes of an induced gene cluster are shown below the reactions. They code for subunits similar to PpcA and PpcD, which are proposed to be involved in benzene carboxylation, and for a proposed benzoate-CoA ligase (Blz). Moreover, a gene coding for an ortholog of UbiX (see above) is located in the gene cluster. (c) Proposed pathway of naphthalene degradation via carboxylation. A gene cluster containing the highly induced nacA gene is shown below the reactions. Whereas NacA is similar to the PpcA subunit of phenylphosphate carboxylase, several potential orthologs of PpcB and PpcC are encoded in close vicinity (indicated as nacB1-3 and nacC, respectively)

Alkene Hydration Anaerobic alkene degradation is proposed to proceed via hydration of the double bound to a primary alcohol, or via subterminal carbon dioxide addition leading to the formation of branched fatty acids (Grossi et al. 2007). The first enzyme involved in anaerobic metabolism of an alkene substrate has

only recently been identified and characterized. It is involved in the degradation pathway of the terpenoid alcohol linalool by the Betaproteobacterium Castellaniella defragrans and catalyzes the reversible dehydration/hydration of linalool to the unsaturated hydrocarbon myrcene as well as the isomerization of linalool to geraniol (> Fig. 17.11), and therefore was named linalool dehydratase/isomerase (LDI) (Brodkorb et al. 2010).

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periplasm

cytoplasm

H2O

H2O myrcene

2 [H] 2 [H] HO

HO OH linalool

geraniol

-OOC

geraniol

unknown further pathway

geranate

. Fig. 17.11 Proposed pathway of linalool degradation. The initial enzyme linalool dehydratase/isomerase catalyzes the reversible dehydration of linalool to myrcene and generation of geraniol. Instead of assuming a direct isomerization of linalool to geraniol (Brodkorb et al. 2010), we suggest the reaction to proceed by subsequent reversible dehydration and hydratation steps, involving the hydrocarbon myrcene as intermediate. Geraniol is further oxidized in two steps to geranate. The reactions involved in further degradation of geranate are still unknown

The mechanism for linalool isomerization to geraniol via myrcene as intermediate shown in > Fig. 17.11 deviates from that assumed by Brodkorb et al. (2010), but equally fits to the available data. The enzyme apparently belongs to a new enzyme family, does not contain any apparent cofactors, and depends in its activity on added thiol reagents like DTT. It appears to be located in the periplasm, being secreted via the Sec system (Brodkorb et al. 2010). The further degradation pathway involves a two-step oxidation of geraniol to geranic acid, but the further reactions of geranic acid degradation are unknown.

‘‘Reverse Methanogenesis’’ by Sulfate-Reducing Consortia Anaerobic oxidation of methane to CO2/HCO3 has been reported for sulfate-reducing consortia and the nitrate-reducing bacterium Methylomirabilis oxyfera (Boetius et al. 2000; Nauhaus et al. 2002; Orphan et al. 2002; Raghoebarsing et al. 2006; Ettwig et al. 2008, 2009). A recent publication also described anaerobic methane oxidation in marine habitats being driven by the reduction of Mn(IV)- and Fe(III)-ions (Beal et al. 2009). Since the ‘‘intra-aerobic’’ species Methylomirabilis oxyfera has been shown to utilize methane monooxygenase for the initial attack on methane with the oxygen cosubstrate developed from nitrite (Ettwig et al. 2010), only the truly anaerobic biochemical mechanisms employed under sulfatereducing and (possibly) metal-ion-reducing conditions will be discussed here. Anaerobic oxidation of methane under sulfate-reducing conditions is known for consortia of Euryarchaeota in

association with sulfate-reducing Deltaproteobacteria. The Archaea belong to novel phylogenetic groups affiliated to Methanosarcinales and are apparently the methane-oxidizing partners of the consortia by reverting the final steps of methanogenesis (Boetius et al. 2000; Knittel et al. 2005). The terminal step of methanogenesis is the reduction of methyl-coenzyme M (CoMSH) with coenzyme B (CoBSH) to form methane and a CoM-S-S-CoB-heterodisulfide (Pelmenschikov et al. 2002) (> Fig. 17.12). This reaction is catalyzed in all methanogens by methyl-coenzyme M reductase (Mcr), which has an (abg)2 composition and contains the nickel-tetrapyrrole cofactor F430 (Hallam et al. 2003; Kru¨ger et al. 2003; Thauer et al. 2008). F430 nickel cofactors and Mcr’s with very similar characteristics as those known from methanogenic Archaea were isolated directly from methanotrophic mats from the Black Sea, which were dominated by Archaea of the ANME-1 group. Remarkably, two abundant nickel-containing Mcr-paralogs were extracted, one of which contained a normal F430 cofactor, whereas the other contained a modified F430 cofactor (17(2)-methylthioF430) (Mayr et al. 2008). The two Mcr isoenzymes were assigned to two different subgroups of anaerobic methanotrophs, and both were shown to be highly expressed. However, these slight differences in Mcr cofactor content are not necessarily indicating the presence of a subpopulation of methanogenic Archaea in the methanotrophic consortia, because the reaction of Mcr has recently been shown to be generally reversible (Scheller et al. 2010b). Therefore, any type of Mcr from methanogenic or methanotrophic Archaea and with any F430 cofactor version appears capable to catalyze both methane formation and the conversion of methane to methyl-CoM

Anaerobic Biodegradation of Hydrocarbons Including Methane

CH4

CH3

H

Ni(III)

Ni(I)

CoM-S-CH3

CoM-S-S-CoB

SO42− [H]

H3C

HS−

CoB-S−

CoM S

H3C

HS-CoM Ni(III)

Ni(I) CO 2

17

CoB-SH

. Fig. 17.12 Proposed initial reaction of anaerobic methane oxidation via ‘‘reverse methanogenesis.’’ Reversible cleavage of one of the C–H bonds of methane may involve formation of a transient complex with the Ni(I)-state of the F430 tetrapyrrole cofactor of methyl-CoM reductase, combined with a conformation change of the F430 to an unstable intermediate with bound methyl and hydride groups. From there, the reaction may then either run backward to methane or forward to a methyl-Ni(III)-adduct, accompanied by stepwise transfer of two electrons to the heterodisulfide of CoM and CoB, yielding the reduced forms of these coenzymes (Modified from Scheller et al. 2010b). The electron transfer process probably occurs via a radical anion of the heterodisulfide and the thiyl-radical form of CoB as intermediates. The F430-bound methyl group is then thought to recombine with reduced CoM to methyl-CoM, reconstituting F430 to the Ni(I)-state. The reaction continues with further oxidation steps of methyl-CoM and the transfer of redox equivalents to the syntrophic sulfate-reducing partner organism at a still unknown step. The assumed fate of the proton cleaved off from methane is highlighted in red to visualize the observed hydrogen/deuterium exchange between methane and methyl-CoM

(Thauer and Shima 2008; Scheller et al. 2010a). The observed differences in F430-cofactor content in methanotrophic mats may then be more related to phylogenetic differences of the archaeal strains than to mechanistic differences of the enzymes involved (Thauer 2011). The new enzymological data on Mcr fit well to older observations of methanogens re-oxidizing a small percentage of the methane produced, albeit apparently without energy gain (Zehnder and Brock 1979; Harder 1997). Therefore, the proposed ‘‘reverse methanogenesis’’ mechanism involved in AOM seems to be biochemically very plausible. The reversibility of the Mcr reaction and recent data on potential conformation shifts of the F430 cofactor during the reaction also raised questions about the reaction mechanism of this enzyme. A direct involvement of the highly reactive Ni(I) redox state of F430 has been proposed for the initial reaction of breaking one of the C-H bonds for methane oxidation, as shown in > Fig. 17.12 (Scheller et al. 2010b). Based on the new data on the mechanism of Mcr, methanotrophic Archaea may differ from normal methanogens not too much in the mechanisms of generating or consuming methane, but mostly in their still unknown mechanisms of transferring redox equivalents to the sulfate-reducing bacterial partner organisms. Recently, a possible transfer mechanism involving the transfer of methyl sulfides between the syntrophic partners was proposed (Moran et al. 2008), but other mechanisms are still not excluded. The genes coding for the

methyl-coenzyme M reductase (Mcr) paralogs and further enzymes likely involved in the methane-oxidation pathway were identified in metagenomic sequences of methanotrophic mats (Hallam et al. 2003, 2004; Meyerdierks et al. 2005). The genes coding for the three subunits of Mcr, mcrBGA, are often forming an operon with two additional genes of unknown functions (mcrC and mcrD). The gene encoding the a-subunit of Mcr, mcrA, has been established as key genetic marker to identify and determine the phylogenetic positions of methanogens as well as anaerobic methanotrophs (Friedrich 2005).

Applications Bioremediation One of the most important applied aspects in studying anaerobic hydrocarbon-degrading microorganisms in the past decade has been bioremediation of contaminated groundwater aquifers. Because of the limited solubility of oxygen in water, heavy contamination of groundwater always results in the development of an anaerobic zone by the metabolic activity of aerobic microorganisms. Therefore, in situ bioremediation in such environments is largely a domain of the anaerobic hydrocarbon degraders, despite their much slower metabolic rates compared to aerobes. Recent developments have been

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the design of analytical tools to monitor the processes going on in anaerobic underground sites, which are based on the identification of the microorganisms present by molecular probes (Beller et al. 2002; Hosoda et al. 2005; Da Silva and Alvarez 2007; Kuntze et al. 2008, 2011; Winderl et al. 2008; Kazy et al. 2010) or via proteomic analysis and stable isotope probing methods (Griebler et al. 2004; Kunapuli et al. 2007; Herrmann et al. 2010; Jehmlich et al. 2010; Winderl et al. 2010). Assessment of the in situ microbial processes is measurable by the identification of key metabolites linked to the metabolic pathways typical for anaerobic hydrocarbon metabolism (Beller 2000; Gieg and Suflita 2002; Reusser et al. 2002; Nijenhuis et al. 2007; Gieg et al. 2009; Parisi et al. 2009; Jobelius et al. 2011) and the analysis of the isotopic signatures of the identified metabolites and the remaining contaminants (Meckenstock et al. 1999b; Vieth et al. 2005; Fischer et al. 2008; Vogt et al. 2008). A more ambitious aim of this line of research is to enhance the rates of biological pollutant degradation by bioaugmentation, physical or chemical treatment of the contaminated sites (Hunkeler et al. 2002; Da Silva and Alvarez 2004; Kasai et al. 2007; Bauer et al. 2009).

Microbial Enhanced Oil Recovery and Biofuels Another recent development is centered around the idea of using microbial activities to enhance the recovery of oil or gas from exhausted or nearly exhausted reservoirs (for recent reviews, see Brown 2010; Gray et al. 2010). The usually used methods involve inoculating chemotrophic surfactant- or gas-producing bacteria of various physiological types (mostly thermophilic aerobic or fermentative bacteria) with or without added nutrients into the oil reservoirs to increase the pressure in the reservoir and/or to decrease the viscosity of the residual heavy oil in order to retrieve more crude oil. First trials have been started to introduce oil-degrading methanogenic consortia into such oil reservoirs, which may degrade the hardly mobile heavy oil compounds into more easily retrievable short-chain alkanes and methane (Gieg et al. 2008, 2010). Moreover, a better understanding of the organisms and biochemical processes involved in anaerobic hydrocarbon degradation will also help in designing new processes of generating hydrocarbons biologically from renewable sources. This knowledge may either be necessary to prevent futile degradation cycles in currently emerging biotechnological hydrocarbon production methodologies, e.g., from fatty acids or isoprenoids (Wackett 2011), or even to incorporate some of the reactions into new synthetic pathways for microbial hydrocarbon production.

Fine Chemicals/Biotechnology Finally, the unusual biochemical pathways used in anaerobic hydrocarbon metabolism provide a wealth of new enzymes and new biochemical principles that may be exploited for the production of fine chemicals. Many of the known

pathways contain highly interesting chiral intermediates, which may be of technical interest, e.g., (R)-benzylsuccinate as monomer for biodegradable plastics (Xu and Guo 2010), or chiral alcohols as building blocks for chemical synthesis. Moreover, many of the enzymes involved have a very broad substrate specificity and catalyze highly interesting enantiomer-specific conversions of many unnatural substrates, which may be of practical interest (Schlieben et al. 2005; Ho¨ffken et al. 2006; Mugford et al. 2008).

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Vogt C, Cyrus E, Herklotz I, Schlosser D, Bahr A, Herrmann S, Richnow HH, Fischer A (2008) Evaluation of toluene degradation pathways by two-dimensional stable isotope fractionation. Environ Sci Technol 42:7793–7800 Vogt C, Kleinsteuber S, Richnow HH (2011) Anaerobic benzene degradation by bacteria. Microb Biotechnol 4:710–724 Wackett LP (2011) Engineering microbes to produce biofuels. Curr Opin Biotechnol 22:388–393 Wardlaw GD, Arey JS, Reddy CM, Nelson RK, Ventura GT, Valentine DL (2008) Disentangling oil weathering at a marine seep using GC  GC: broad metabolic specificity accompanies subsurface petroleum biodegradation. Environ Sci Technol 42:7166–7173 Weelink SAB, Tan NCG, ten Broeke H, van den Kieboom C, van Doesburg W, Langenhoff AAM, Gerritse J, Junca H, Stams AJM (2008) Isolation and characterization of Alicycliphilus denitrificans strain BC, which grows on benzene with chlorate as the electron acceptor. Appl Environ Microbiol 74:6672–6681 Weelink SA, van Doesburg W, Saia FT, Rijpstra WI, Ro¨ling WF, Smidt H, Stams AJ (2009) A strictly anaerobic betaproteobacterium Georgfuchsia toluolica gen. nov., sp. nov. degrades aromatic compounds with Fe(III), Mn(IV) or nitrate as an electron acceptor. FEMS Microbiol Ecol 70:575–585 Widdel F, Bak F (1992) Gram-negative mesophilic sulfate-reducing bacteria. In: Balows A, Tru¨per HG, Dworkin W, Harder W, Schleifer K-H (eds) The prokaryotes, 2nd edn. Springer, Berlin, pp 3352–3378 Widdel F, Grundmann O (2010) Biochemistry of the anaerobic degradation of non-methane alkanes. In: Timmis KN (ed) Handbook of hydrocarbon and lipid microbiology. Springer, Berlin, pp 909–924 Widdel F, Musat F (2010) Diversity and common principles in enzymatic activation of hydrocarbons. In: Timmis KN (ed) Handbook of hydrocarbon and lipid microbiology. Springer, Berlin, pp 983–1009 Widdel F, Rabus R (2001) Anaerobic biodegradation of saturated and aromatic hydrocarbons. Curr Opin Biotechnol 12:259–276 Wilkes H, Schwarzbauer J (2010) Hydrocarbons: an introduction to structure, physico-chemical properties and natural occurrence. In: Timmis KN (ed) Handbook of hydrocarbon and lipid microbiology. Springer, Berlin, pp 5–48 Wilkes H, Boreham C, Harms G, Zengler K, Rabus R (2000) Anaerobic degradation and carbon isotopic fractionation of alkylbenzenes in crude oil by sulphate-reducing bacteria. Org Geochem 31:101–115 Wilkes H, Rabus R, Fischer T, Armstroff A, Behrends A, Widdel F (2002) Anaerobic degradation of n-hexane in a denitrifying bacterium: further degradation of the initial intermediate (1-methylpentyl)succinate via C-skeleton rearrangement. Arch Microbiol 177:235–243 Wilkes H, Ku¨hner S, Bolm C, Fischer T, Classen A, Widdel F, Rabus R (2003) Formation of n-alkane- and cycloalkane-derived organic acids during

anaerobic growth of a denitrifying bacterium with crude oil. Org Geochem 34:1313–1323 Wilkinson ER (1972) California offshore oil and gas seeps. California Division of Oil and Gas, Sacramento Winderl C, Anneser B, Griebler C, Meckenstock RU, Lueders T (2008) Depthresolved quantification of anaerobic toluene degraders and aquifer microbial community patterns in distinct redox zones of a tar oil contaminant plume. Appl Environ Microbiol 74:792–801 Winderl C, Penning H, von Netzer F, Meckenstock RU, Lueders T (2010) DNASIP identifies sulfate-reducing Clostridia as important toluene degraders in tar-oil-contaminated aquifer sediment. ISME J 4:1314–1325 Wolterink A, Schiltz E, Hagedoorn PL, Hagen WR, Kengen SWM, Stams AJM (2003) Characterization of the chlorate reductase from Pseudomonas chloritidismutans. J Bacteriol 185:3210–3213 Wu ML, Ettwig KF, Jetten MSM, Strous M, Keltjens JT, van Niftrik L (2011) A new intra-aerobic metabolism in the nitrite-dependent anaerobic methaneoxidizing bacterium Candidatus ‘Methylomirabilis oxyfera’. Biochem Soc Trans 39:243–248 Xu J, Guo BH (2010) Poly(butylene succinate) and its copolymers: research, development and industrialization. Biotechnol J 5:1149–1163 Yagi JM, Suflita JM, Gieg LM, DeRito CM, Jeon CO, Madsen EL (2010) Subsurface cycling of nitrogen and anaerobic aromatic hydrocarbon biodegradation revealed by nucleic acid and metabolic biomarkers. Appl Environ Microbiol 76:3124–3134 Zedelius J, Rabus R, Grundmann O, Werner I, Brodkorb D, Schreiber F, Ehrenreich P, Behrends A, Wilkes H, Kube M, Reinhardt R, Widdel F (2011) Alkane degradation under anoxic conditions by a nitrate-reducing bacterium with possible involvement of the electron acceptor in substrate activation. Environ Microbiol Rep 3:125–135 Zehnder AJB, Brock TD (1979) Methane formation and methane oxidation by methanogenic bacteria. J Bacteriol 137:420–432 Zengler K, Heider J, Rossello-Mora R, Widdel F (1999a) Phototrophic utilization of toluene under anoxic conditions by a new strain of Blastochloris sulfoviridis. Arch Microbiol 172:204–212 Zengler K, Richnow HH, Rossello-Mora R, Michaelis W, Widdel F (1999b) Methane formation from long-chain alkanes by anaerobic microorganisms. Nature 401:266–269 Zhang XM, Young LY (1997) Carboxylation as an initial reaction in the anaerobic metabolism of naphthalene and phenanthrene by sulfidogenic consortia. Appl Environ Microbiol 63:4759–4764 Zhou JZ, Fries MR, Cheesanford JC, Tiedje JM (1995) Phylogenetic analyses of a new group of denitrifiers capable of anaerobic growth on toluene and description of Azoarcus tolulyticus sp. nov. Int J Syst Bacteriol 45:500–506

18 Physiology and Biochemistry of the Methane-Producing Archaea Reiner Hedderich1 . William B. Whitman2 1 Max Planck Institute fu¨r Terrestriche, Mikrobiologie, Marburg, Germany 2 Department of Microbiology, University of Georgia, Athens, GA, USA

Ecology of Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 636 Biogeochemistry of Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . 637 Systematics of the Methanoarchaea . . . . . . . . . . . . . . . . . . . . . . . 637 Pathways of Methanogenesis: An Overview . . . . . . . . . . . . . . . 640 Key Reactions in Biological Methane Formation . . . . . . . . . . 642 The Final Step of Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . 642 Energy Conservation via Disulfide Respiration . . . . . . . . . 642 Reductive Activation of CO2 to Formylmethanofuran . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 645 Tetrahydromethanopterin Replaces Tetrahydrofolate in the C1 Pathway of Methanoarchaea . . . . . . . . . . . . . . . . . . 646 A Sodium Ion Pumping Methyltransferase . . . . . . . . . . . . . 646 Activation of Methanol and Methylamines . . . . . . . . . . . . . 649 The Aceticlastic Reaction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 649 The Hydrogenases of Methanoarchaea: A Summary . . . . . . 650 F420-Reducing Hydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 651 H2-Forming Methylene-H4MPT Dehydrogenase . . . . . . . 651 F420-Nonreducing Hydrogenase . . . . . . . . . . . . . . . . . . . . . . . . . 652 Methanophenazine-Reducing Hydrogenases . . . . . . . . . . . . 652 Energy-Converting [NiFe] Hydrogenases . . . . . . . . . . . . . . . 652 Methanogenic Coenzymes and Enzymes in Nonmethanogenic Archaea and Bacteria . . . . . . . . . . . . . . . . . . 653 Sulfate-Reducing Archaea Use Three Methanogenic Coenzymes for the Oxidation of Reduced C1 Compounds to CO2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 653 Tetrahydromethanopterin-Dependent Formaldehyde Oxidation in Methylotrophic Bacteria . . . . . . . . . . . . . . . . . . 653 F420 in Nonmethanogenic Organisms . . . . . . . . . . . . . . . . . . . 654 CoM-SH in Bacterial Aliphatic Epoxide Carboxylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 654 Do Anaerobic Methane Oxidizers Use the Methanogenic Pathway in Reverse? . . . . . . . . . . . . . . . . . . . . . 654 Regulation of Gene Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . 654 Regulation of Catabolic Enzymes by Substrate Availability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 654 Regulation of Catabolic Enzymes by Trace Element Availability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 655 Regulation of Nitrogen Assimilation . . . . . . . . . . . . . . . . . . . . 656

Bioenergetics of Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 656 Coupling Sites in Methanogenesis . . . . . . . . . . . . . . . . . . . . . . 656 Growth Yields . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 657 Genomes of the Methanoarchaea . . . . . . . . . . . . . . . . . . . . . . . 658 Abstract Methanoarchaea are an ancient monophyletic lineage within the Euryarchaeota, thriving by chemolithotrophic energy metabolism. The chapter discusses three major problems: (i) distinguishing taxa on the basis of their similar phenotype; (ii) the extreme genetic diversity of methanogens and, consequently, the high variability of their metabolism; and (iii) based upon analysis of environmental DNA, the notion that the cultured methanogens represent a very sparse sampling of the likely diversity in nature. Hence, knowledge of the species richness and metabolic diversity of this group is necessarily incomplete. The following will focus on the large differences in cellular structure, metabolic pathways, and regulation. Methanoarchaea derive their metabolic energy from the conversion of a restricted number of substrates to methane. Most methanoarchaea can reduce CO2 to CH4. The major electron donors for this reduction are H2 and formate. In addition, some methanoarchaea can use alcohols like 2-propanol, 2-butanol, cyclopentanol, and ethanol as electron donors. The second type of substrate for methanogenesis includes C-1 compounds containing a methyl group carbon bonded to O, N, or S. Compounds of this type include methanol, monomethylamine, dimethylamine, trimethylamine, tetramethylammonium, dimethylsulfide, and methane thiol. The third type of substrate is acetate. In this reaction, the methyl (C-2) carbon of acetate is reduced to methane using electrons obtained from the oxidation of the carboxyl (C-1) carbon of acetate. This reaction is called the ’aceticlastic reaction’ because it results in the splitting of acetate into methane and CO2. Key reactions of the different methanogenic pathways including, electron chains and structures, as well as genomic information and methanogenic coenzymes and enzymes in nonmethanogenic Archaea and Bacteria are described. The methane-producing archaea or methanoarchaea are distinguished by their ability to obtain all or most of their energy for growth from the process of methane biosynthesis or methanogenesis. To date, no methanogens have been identified that can grow without producing methane, and these archaea are

E. Rosenberg et al. (eds.), The Prokaryotes – Prokaryotic Physiology and Biochemistry, DOI 10.1007/978-3-642-30141-4_81, # Springer-Verlag Berlin Heidelberg 2013

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Physiology and Biochemistry of the Methane-Producing Archaea

Methanogenium

Methanomicr

s

Methanospirillum Methanosaeta

Methanosar

Methanosarcina Haloferax Thermoplasma Ferroglobus Archaeoglobus Methanothermus Methanothermobacter Methanococcus Methanocaidococcus Pyrococcus Methanopyrus

Methanop

0.05

. Fig. 18.1 Phylogeny of the methanoarchaea and related euryarchaeota. The gene tree of the 16S rRNA was calculated with the Fitch-Margoliash algorithm in PHYLIP and about 1,260 bp of the small subunit rRNA sequences for representatives of each genus. The orders of methanogens are indicated on the right. The scale bar represents the Jukes-Cantor evolutionary distance

all obligate methane producers that are uniquely specialized for this lifestyle. Methanogenesis is an anaerobic respiration, but its complexity and commitment of resources far exceed that found in other common respiratory processes. For instance, it requires the biosynthesis of six unusual coenzymes; a long, multistep pathway for methane; and a number of unique membranebound enzyme complexes for coupling to the proton motive force (see below). Given the complexity of this process, it is not surprising that methanogens appear to be monophyletic. Hence, all modern methanoarchaea possess an ancient ancestor within the Euryarchaeota (> Fig. 18.1). Although the branching order of the deep branches is not certain, several lineages of nonmethanogenic archaea appear within this clade, which argues for the antiquity of the methanogen group. Thus, the phylogeny suggests that the lineages represented by Haloferax, Thermoplasma, and Archaeoglobus were derived from methanogenic ancestors. The sulfate-reducer Archaeoglobus also possesses many of the unusual coenzymes found in methanogens, providing further evidence of this hypothesis. The methanoarchaea are also the only archaea currently cultivated that are truly cosmopolitan, being found in a wide variety of the anaerobic environments on earth. Thus, in addition to many temperate habitats, methanogens are also common at extremes of temperature and salinity. As examples, psychrophilic and psychrotolerant species isolated from a meromictic lake in Antarctica grow well at 0–5  C (Franzmann et al. 1992, 1997), and hyperthermophilic species obtained from geothermal springs and submarine vents grow well up to 110  C (e.g., Jones et al. 1983; Kurr et al. 1991). While many methanogens grow well in freshwater, the extremely halophilic Methanohalobium evestigatus requires 4.3 M NaCl for optimal

growth (Zhilina and Zavarzin 1987). The broad distribution of the methanogens is in contrast to the other cultivated archaea, which appear to be limited to extreme environments of high temperature, low pH, or high salinity. Presumably, archaea fair poorly in direct competition with temperate bacteria and eukaryotes and are limited to either extreme habitats or niches where they do not compete with bacteria or eukaryotes. This hypothesis implies that the currently uncultivated Crenarchaeota common in temperate soil and marine environments occupy a physiological niche unavailable to the bacteria. It also poses an interesting physiological mystery as to why bacteria might outcompete archaea in many temperate environments.

Ecology of Methanogenesis While the ecology of the methanoarchaea is discussed in detail in the chapters about specific groups, some generalities are appropriate here (for a more detailed review, see Zinder et al. 1993). The methanoarchaea flourish in anaerobic environments where sulfate, oxidized metals, and nitrate are absent. In these environments, the substrates for methanogenesis are readily available as the fermentation products of bacteria and eukaryotes, and the methanoarchaea catalyze the terminal step in the anaerobic food chain where complex polymers are converted to methane and CO2. In this food chain, polymers are first degraded by specialized microorganisms, like the cellulolytic bacteria, to produce simple sugars (such as glucose), disaccharides (such as cellobiose), lactate, volatile fatty acids (VFAs; such as acetate, propionate, and butyrate), and alcohols (such as ethanol). These products are further metabolized by the

Physiology and Biochemistry of the Methane-Producing Archaea

intermetabolic group. These microorganisms convert simple sugars to VFAs and alcohols. They also convert VFAs and alcohols to acetate, H2, and CO2, which are major substrates for the methanoarchaea. Molecular hydrogen (H2) is a key intermediate in this process. Under standard conditions, when the H2 partial pressure is 1 atm., the fermentations of VFAs and alcohols to acetate and H2 are thermodynamically unfavorable. Therefore, microorganisms that catalyze these reactions cannot grow, and toxic levels of the VFAs accumulate. However, if the methanoarchaea are present, H2 is rapidly metabolized, and its partial pressure is maintained below 10 3–10 4 atmospheres. Under these conditions, the fermentations of VFAs and alcohols are thermodynamically favorable, they are rapidly metabolized, and their concentrations are maintained below the toxic levels. This interaction between the H2-producing intermetabolic organisms and the H2-consuming methanogens is an example of interspecies hydrogen transfer (Chap. 21, ‘‘Syntrophism Among Prokaryotes’’ in this Vol. 2). In many anaerobic environments, it is a key regulatory mechanism. Because the H2-consuming methanogens play a critical role, they are said to ‘‘pull’’ the fermentation of complex organic polymers to methane and CO2. Interspecies hydrogen transfer is not limited to methanogenic food chains. In environments rich in sulfate, oxidized metals, or nitrate, anaerobic bacteria oxidize H2, VFAs, and alcohols. When sulfate is present, this activity is catalyzed by the sulfate-reducing bacteria. Because the oxidation of H2 with sulfate as the electron acceptor is thermodynamically more favorable than when CO2 is the electron acceptor (as in methanogenesis), the sulfate-reducing bacteria outcompete the methanogens for H2. For similar reasons, the sulfate-reducing bacteria also outcompete the methanogens for other important substrates like acetate and formate. Therefore, methanogenesis is greatly limited in marine sediments that are rich in sulfate. The oxidation of H2 with nitrate, Fe+3, and Mn+4 as electron acceptors is also thermodynamically more favorable than methanogenesis. The denitrifying and iron- and magnesiumreducing bacteria also outcompete the methanogens when these electron acceptors are present. In conclusion, methanogenesis only dominates in habitats where CO2 is the only abundant electron acceptor for anaerobic respiration. Just as aerobic microorganisms rapidly deplete the O2 in environments rich in organic matter to establish anaerobic conditions, sulfate-reducing bacteria, iron- and magnesiumreducing bacteria, and denitrifying bacteria frequently consume all the sulfate, Fe+3, Mn+4, and nitrate in anaerobic environments and rapidly establish the conditions for methanogenesis. In these environments, CO2 is seldom limiting because it is also a major fermentation product. Thus, methanogenesis is the dominant process in many anaerobic environments that contain large amounts of easily degradable organic matter. Especially important environments of this type include freshwater sediments found in lakes, ponds, marshes, and rice paddies. Although methanoarchaea are most frequently found at the bottom of anaerobic food chains associated with the intermetabolic microorganisms, in some ecosystems they are the primary

18

consumers of geochemically produced H2 and CO2. The submarine hydrothermal vents found on the ocean floor expel large volumes of very hot water containing H2 and H2S. As the water cools from several hundred degrees Celsius to the temperature of the ocean, zones suitable for the growth of thermophilic methanoarchaea are established. The methane produced escapes into the surrounding water where it is utilized by symbiotic methylotrophic bacteria living in marine invertebrates. In this ecosystem, the methanogens are at the top of the food chain.

Biogeochemistry of Methanogenesis Methane is a major trace gas found in the earth’s atmosphere, and most of the atmospheric methane is produced by the methanoarchaea (Conrad 1996; Monson and Holland 2001; Reeburo¨gh 2003). In the last several hundred years, the atmospheric concentration has more than doubled, to reach about 1.8 ppm in 1998. This increase in methane concentration is responsible for about 20 % of the increased greenhouse effect observed for all radiatively important trace gases. Currently, methane emissions to the atmosphere are about 500–600 teragrams (Tg) of CH4 year 1. Once in the atmosphere, methane has a half-life of about 8.4 years, being removed primarily by reactions with hydroxyl radicals in the troposphere. Major sources of atmospheric methane are shown in > Table 18.1. The most important biogenic emissions are from habitats where gases formed anaerobically can exchange rapidly with the atmosphere, such as wetlands and rice paddies. Similarly, large amounts of methane are emitted from the rumen of livestock, which escapes directly to the atmosphere by eructation. However, large amounts of methane, probably about 700 Tg of CH4 year 1, are also consumed by aerobic and anaerobic methane oxidizers without ever appearing in the atmosphere (Reeburgh et al. 1993; Valentine and Reeburgh 2000; Reeburo¨gh 2003). Rice paddies are particularly active sites for methane oxidation, where oxygen is available in the root zone owing to transport through the plants. This allows for growth of the aerobic methane-oxidizing bacteria and consumption of 45–90 % of the methane produced. Given that about 80 % of the methane emitted to the atmosphere and nearly all of the methane oxidized prior to release to the atmosphere is microbially produced, an estimate of total microbial methane production is about 1,100 Tg of CH4 year 1, which represents about 825 Tg of C year 1. During the production of methane from carbohydrates, one mole of CO2 is formed for every mole of CH4. Thus, the total carbon processed in these systems is about 1,650 Tg of C year 1, or 1.65 % of the C fixed by photosynthesis each year (Schlesinger 1991). This estimate emphasizes the biogeochemical significance of this link in the carbon cycle.

Systematics of the Methanoarchaea Phylogenetic analyses of the rRNA and other genes indicate that the methanoarchaea are an ancient monophyletic lineage within

637

638

18

Physiology and Biochemistry of the Methane-Producing Archaea

. Table 18.1 Sources of atmospheric methane Sources of methane

Methane evolved (Tg of CH4/year)

Biogenic sources Natural wetlands

92

Rice paddies

88

Livestock

81

Manure decomposition

14

Termites

25–150

Landfills

15–81

Oceans

38–308

Tundra

42

Subtotal

395–856

Other sources Biomass burning

50

Coal mining

10–35

Venting and flaring

15–30

Industrial and pipeline losses 15–45 Methane hydrates

5

Subtotal

95–165

Totala

600 (490–1,021)

Modified from Tyler (1991) and Reeburo¨gh (2003) Abbreviation: Tg, teragram, 1012 g or one million metric tons a Best estimate with range in parentheses

the Euryarchaeota (> Fig. 18.1). Although the branching orders of the deep groups of the Euryarchaeota are not known for certain, a number of nonmethanogenic organisms, such as the halobacteria and the sulfate-reducing Archaeoglobus, appear to have arisen within the methanogenic lineage. To produce descendants with very different phenotypic properties, the methanogenic lineage must be very ancient. The current taxonomy for methanogens follows the general schema of Boone et al. (1993) and Whitman et al. (2001), which tried to form taxa of similar phylogenetic depth among groups of fairly unrelated organisms. This work attempted to deal with three major problems. First, because of their chemolithotrophic energy metabolism, it is often difficult to distinguish taxa on the basis of phenotype. Second, in spite of the phenotypic similarity, the methanogens are genetically, extremely diverse. The high genetic diversity suggests that even though many of these organisms appear to do the same thing, the way they do these things are very different. Detailed studies of the physiology have tended to support this view, and large differences in cellular structure, metabolic pathways, and regulation have been observed. And third, on the basis of ribosomal RNA gene libraries of environmental DNA, many more taxa await to be isolated and characterized (see below). The cultured methanogens represent a very sparse sampling of the likely diversity in nature, and our knowledge of this group is necessarily incomplete. Thus, the current taxonomy

is best considered a work in progress with plenty of opportunities for improvement. The methanoarchaea are a diverse group of organisms, containing five well-established orders and 31 genera (> Table 18.2). For species, less than 70 % DNA hybridization of the genomic DNAs is considered definitive evidence for novel species (Wayne et al. 1987). In the absence of DNA hybridization data, ribosomal RNA sequence similarity of less than 98 % is considered equivalent evidence for novel species, even though this value is probably very conservative (Stackebrandt and Goebels 1994; Keswani and Whitman 2001). Less than 93–95 % ribosomal RNA sequence similarity is evidence of novel genera, and less than 88–93 % ribosomal RNA sequence similarity is evidence of novel families. The rank of order is then used to recognize deeper phylogenetic differences. In general, this taxonomy is supported by the chemotaxonomy of cellular lipids and distribution of other biological properties. Subsequently, three classes were also proposed: the Methanobacteria (to include the Methanobacteriales), the Methanococci (to include the Methanococcales, Methanomicrobiales, and Methanosarcinales), and the Methanopyri (to include the Methanopyrales; Boone 2001). These classes were inferred from the deep phylogenetic relationships in the ribosomal RNA gene tree, which are imperfectly understood, and their biological significance remains to be further elucidated. A summary of the major genera of methanoarchaea is given in > Table 18.2. In prokaryotic nomenclature, the names of genera of methanoarchaea contain the prefix ‘‘methano-.’’ This prefix distinguishes them from an unrelated group of aerobic bacteria, the methylotrophic bacteria, which consume methane and whose names contain the prefix ‘‘methylo-.’’ More complete descriptions of the nutrition, growth properties, morphology, ecology, and other general properties of these taxa are reviewed in other chapters of this volume as well as Bergey’s Manual of Systematic Bacteriology (Whitman et al. 2001). The descriptions of Methanomethylovorans and Methanomicrococcus can be found in Lomans et al. (1999) and Sprenger et al. (2000). Recent studies of the genus Methanobrevibacter serve to illustrate the incomplete nature of the current systematics of methanogens (Miller 2001). This taxon is abundant in the gastrointestinal tracts of mammals, birds and termites as well as other habitats. It is particularly interesting because it illustrates the diversity of these organisms and some of the potential complexities of their lifestyle. In humans, methane is formed by the methanoarchaea in the anaerobic microflora of the large bowel. About one-third of healthy adults excrete methane gas. Some methane is also absorbed in the blood and excreted from the lungs. The most numerous methanogen in humans is Methanobrevibacter smithii. In people who excrete methane, it is found in numbers of 107–1010 cells per gram dry weight of feces, or between 0.001 % and 12 % of the total number of viable anaerobic prokaryotes (Miller and Wolin 1982). Why the numbers of M. smithii fluctuates so greatly in apparently healthy individuals remains a mystery. Other species of Methanobrevibacter are encountered in the feces of other animals.

Physiology and Biochemistry of the Methane-Producing Archaea

18

. Table 18.2 Taxonomy of the methane-producing archaeaa Morphology

Major energy substratesb

Temperature optimum ( C) Cell wallc

Genus Methanobacterium

Rod

H2, (formate, alcohols)

37–45

Pseudomurein

Methanothermobacter

Rod

H2, (formate)

55–65

Pseudomurein

Methanobrevibacter

Short rod

H2, (formate)

37–40

Pseudomurein

Coccus

H2 + methanol

37

Pseudomurein

Rod

H2

80–88

Pseudomurein + protein

Genus Methanococcus

Coccus

H2, formate

35–40

Protein

Methanothermococcus

Coccus

H2, formate

60–65

Protein

Genus Methanocaldococcus

Coccus

H2

80–85

Protein

Methanotorris

Coccus

H2

88

Protein

Order, family, and genus Order Methanobacteriales Family Methanobacteriaceae

Methanosphaer

a

Family Methanothermaceae Genus Methanothermus Order Methanococcales Family Methanococcaceae

Family Methanocaldococcaceae

Order Methanomicrobiales Family Methanomicrobiaceae Genus Methanomicrobium

Rod

H2, formate

40

Protein

Methanoculleus

Irregular coccus

H2, formate (alcohols)

20–55

Glycoprotein

Methanofollis

Irregular coccus

H2, formate (alcohols)

37–40

Glycoprotein

Methanogenium

Irregular coccus

H2, formate (alcohols)

15–57

Protein

Methanolactnia

Rod

H2 (alcohols)

40

Glycoprotein

Methanoplama

Plate or disc

H2, formate (alcohols)

32–40

Glycoprotein

Spirillum

H2, formate (alcohols)

30–37

Protein + sheath

Family Methanospirillaceae Methanospirillum Family Methanocorpusculaceae Genus Methanocorpusculum

Small coccus

H2, formate (alcohols)

30–40

Glycoprotein

Methanocalculusd

Irregular coccus

H2, formate

30–40

ND

Genus Methanosarcina

Coccus, packets

Methanol, MeNH2, (H2, Ac, DMS)

35–60

Protein + HPS

Methanococcoides

Coccus

Methanol, MeNH2

23–35

Protein

Methanohalophilus

Irregular coccus

Methanol, MeNH2

35–40

Protein

Methanohalobium

Flat polygons

Methanol, MeNH2

40–55

ND

Methanolobus

Irregular coccus

Methanol, MeNH2 (DMS)

37

Glycoprotein

Methanomethylovorans

Coccus, packets

Methanol, MeNH2 DMS, MT

34–37

ND

Methanomicrococcus

Flat polygons

H2 + Methanol, H2 + MeNH2

39

ND

Order Methanosarcinales Family Methanosarcinaceae

639

640

18

Physiology and Biochemistry of the Methane-Producing Archaea

. Table 18.2 (continued) Order, family, and genus

Morphology

Major energy substratesb

Temperature optimum ( C) Cell wallc

Methanosalsum

Irregular coccus

Methanol, MeH2, DMS

35–45

ND

Rod

Ac

35–60

Protein + sheath

Rod

H2

98

Pseudomurein

Family Methanosaetaceae Genus Methanosaeta (Methanothrix) Order Methanopyrales Family Methanopyraceae Genus Methanopyrus

Abbreviations: MeNH2, methylamines (monomethylamine, dimethylamine, and trimethylamine); DMS, dimethylsulfide; MT, methanethiol; Ac is acetate; HPS, heteropolysaccharide; ND, not determined a All of the methanoarchaea are members of the phylum Euryarchaeota b Major energy substrates for methane synthesis. Alcohols are some or all of ethanol, isopropanol, isobutanol, and cyclopentanol. Parentheses means utilized by some but not all species or strains c Cell wall components d Placement in higher taxon is tentative

Methanobrevibacter gottschalkii was isolated from horse and pig feces, Methanobrevibacter thaueri was isolated from cattle feces, Methanobrevibacter wolinii was isolated from sheep feces, and Methanobrevibacter woesei was isolated from goose feces (Miller and Lin 2002). These characterized organisms apparently only represent a small fraction of the diversity present in nature. The rumen is another major habitat for methanogens, and about 10–20 % of the total methane emitted to the earth’s atmosphere originates in the rumen of cows, sheep, and other mammals. In this habitat, complex polymers from grass and other forages are degraded to acetate, volatile fatty acids, H2, and CO2 by the cellulolytic and intermetabolic groups of bacteria, fungi, and protozoans (Miller 1992). The acetate and VFAs are absorbed by the animal and are major energy sources. Thus, little methane is produced from acetate. The H2 is used to reduce CO2 to methane, which is emitted. Methanogenesis represents a significant energy loss to the animal, and up to 10 % of the caloric content of the feed may be lost as methane. Methanobrevibacter species are also common in the rumen. For the bovine rumen, Methanobrevibacter ruminantium is the predominant methanogen isolated. However, in a survey of rRNA genes in the rumen of sheep fed different diets, 62 phylotypes of Methanobrevibacter were recognized, many of which were not closely related to described species and were likely to represent members of at least four novel species (Wright et al. 2004). These results demonstrate the incomplete characterization of the methanoarchaea, even from a fairly well-studied environment, and the substantial intraspecies diversity within these organisms. Similarly, an extensive analysis of 120 Methanobrevibacter sequences from cultures and clone libraries suggested the presence of at least ten deep lineages, many of which contained more than one described species (Dighe et al. 2004). Thus, this genus appears to be very deep, with a large amount of interspecies variation as well. Recognizing that only

. Table 18.3 Free energies for typical methanogenic reactions Reaction

GE (kJ/mol of CH4)

Type 1 CO2 + 4H2 ! CH4 + 2H2O

130

4HCOOH ! CH4 + 3CO2 + 2H2O

120

CO2 + 4 (isopropanol) ! CH4 + 4 (acetone) + 2H2O

37

Type 2 CH3OH + H2 ! CH4 + H2O

113

4CH3OH ! 3CH4 + CO2 + 2H2O

103

4CH3NH3Cl + 2H2O ! 3CH4 + CO2 + 4NH4Cl

74

2(CH3)2S + 2H2O ! 3CH4 + CO2 + 2H2S

49

Type 3 CH3COOH ! CH4 + CO2

33

a small fraction of the organisms in even a single genus are in culture leads to the conclusion that the full extent of the diversity of these organisms is largely unknown.

Pathways of Methanogenesis: An Overview Methanoarchaea derive their metabolic energy from the conversion of a restricted number of substrates to methane (> Table 18.3; > Fig. 18.2). Most methanoarchaea can reduce CO2 to CH4. The major electron donors for this reduction are H2 and formate. In addition, some methanoarchaea can use alcohols like 2-propanol, 2-butanol, cyclopentanol, and ethanol as electron donors. For the secondary alcohols, a two-electron

Physiology and Biochemistry of the Methane-Producing Archaea

b

a

CO2

CHO-MFR CHO-H4MPT CH=H4MPT+ H2

CO2 Fdred

F420H2

Fdred CH3-H4MPT

H2

CHO-H4MPT CH=H4MPT+ F420H2

CoM-SH CH3-S-CoM H2

CoB-SH

F420H2

CoM-S-S-CoB

CH4

?

CH2=H4MPT F420H2

Fdred

CH3-H4MPT

CH3-H4MPT CoM-SH 2H+ H2

CH3-S-CoM

CH3-OH

CH3-S-CoM

CoB-SH

CoM-SH

H2

CoM-S-S-CoB

CH4

H2

Fdox CHO-MFR

CH3-CO-S-CoA CO2

CH2=H4MPT H2

c

CH3COOH

ATP

Fd

H2

18

CoB-SH

F420H2 Fdred

CoM-S-S-CoB

?

CH4

. Fig. 18.2 Scheme of methanogenesis from H2/CO2 (a), acetate (b), and methanol (c). Methyl-coenzyme M (CH3-S-CoM) is a central intermediate in all three pathways. It is converted to methane and the heterodisulfide of coenzyme M and coenzyme B (CoM-S-S-CoB). CoM-S-S-CoB thus generated functions as the terminal electron acceptor of different respiratory chains. H2 and reduced coenzyme F420(F420H2) have been identified as electron donors for the reduction of CoM-S-S-CoB. The unknown mechanism of electron transfer from the reduced ferredoxin (Fd(red) ) to CoM-S-S-CoB in acetate and methanol metabolism is symbolized by a question mark. The role of H2 as an intermediate of this reaction is discussed below (see > Fig. 18.14). Abbreviations: CHO-FMR, N-formylmethanofuran; CHO-H4MPT, N5-formyltetrahydromethanopterin; CH = H4MPT+, N5,N10-methenyl-tetrahydromethanopterin; CH2 = H4MPT, N5,N10-methylenetetrahydromethanopterin; and CH3-H4MPT, N5-methyl-tetrahydromethanopterin. For structures of the coenzymes, see > Figs. 18.3, > 18.7, and > 18.9. For simplicity, only tetrahydromethanopterin (H4MPT) is shown. For other methanopterin derivatives, see > Fig. 18.9

oxidation to the ketone is performed. For instance, 2-propanol is oxidized to acetone. Ethanol is somewhat different in that a four-electron oxidation to acetate is performed. Because eight electrons are required to reduce CO2 to methane, four molecules of H2, formate, or 2-propanol are consumed. Even though formate is a reduced C1 compound, it is oxidized to CO2 before reduction to methane. The reduction of CO2 to CH4 proceeds via carrier-bound one-carbon intermediates. Methanofuran (MFR), tetrahydromethanopterin (H4MPT) or its derivatives, and 2-mercaptoethanesulfonate (coenzyme M, CoM-SH) are the three carriers involved (DiMarco et al. 1990; Gorris and van der Drift 1994). These coenzymes were until recently thought to be unique for methanoarchaea but have now also been detected in nonmethanogenic bacteria and archaea (see below). The reaction sequence starts with a two-electron reduction of CO2 and MFR to formyl-MFR where the formyl group is bound to the amino group of the coenzyme. The formyl group is then transferred to the N5 of H4MPT, the formyl-H4MPT thus generated cyclizes to the methenyl-H4MPT, which is reduced in two steps to the methyl-H4MPT. Finally, the methyl group is transferred to the thiol group of coenzyme M. The methyl-thioether formed is reduced to CH4 in the final step of the pathway. The second type of substrate for methanogenesis includes C1 compounds containing a methyl group carbon bonded to O, N, or S. Compounds of this type include methanol,

monomethylamine, dimethylamine, trimethylamine, tetramethylammonium, dimethylsulfide, and methane thiol. The methyl group enters the C1 pathway at the level of coenzyme M and is reduced to methane. The electrons for this reduction are obtained from the oxidation of an additional methyl group to CO2 using the reverse of the steps of the reductive C1 pathway. Because six electrons can be obtained from this oxidation and only two are required to reduce a methyl group to methane, the stoichiometry of this reaction is three molecules of methane formed for every molecule of CO2 formed. In the presence of both a methyl group donor and H2, the methyl oxidation is inhibited and the methyl groups are completely reduced to CH4. An exception to this behavior is found in Methanosphaera and Methanomicrococcus, which lack the ability to oxidize methyl groups. These organisms only grow on methyl compounds when H2 is also present. They are highly specialized for this activity and are unable to reduce CO2 with H2 or other electron donors. The third type of substrate is acetate. In this reaction, the methyl (C-2) carbon of acetate is reduced to methane using electrons obtained from the oxidation of the carboxyl (C-1) carbon of acetate. This reaction is called the ‘‘aceticlastic reaction’’ because it results in the splitting of acetate into methane and CO2. In this metabolism, the methyl group enters the C1 pathway at the level of methyl-H4MPT.

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Key reactions of the different methanogenic pathways will be described below. For a detailed description of methanogenesis, the reader is referred to reviews (Thauer 1998; Deppenmeier et al. 1999; Ferry 1999; Deppenmeier 2002). For a historical overview on methanogenesis, see Wolfe (1991).

Key Reactions in Biological Methane Formation

conservation and their location near the active site suggest an important function of these residues for the formation of the active site and catalysis. The reaction catalyzed by Mcr is rather unusual. Although the actual mechanism is still elusive, all of the proposed catalytic mechanisms involve radical chemistry (Goenrich et al. 2004). The closed hydrophobic environment of the substrate-binding pocket may be optimal for a mechanism employing unstable free radicals.

The Final Step of Methanogenesis

Energy Conservation via Disulfide Respiration

Although every pathway starts out differently, they all end with the same step, the reaction of methyl-coenzyme M (CH3-S-CoM) with a second thiol coenzyme, called ‘‘coenzyme B’’ (CoB-SH), to form methane and the mixed disulfide (also called ‘‘heterodisulfide,’’ CoM-S-S-CoB) of coenzyme M and coenzyme B (> Fig. 18.3). This reaction is catalyzed by methylcoenzyme M reductase (Mcr), making Mcr the key enzyme in methanogenesis (Thauer 1998). In its active site, this enzyme contains a unique prosthetic group, which is a nickel (Ni) porphinoid called ‘‘coenzyme F430’’ (> Fig. 18.3). For the enzyme to be active, Ni has to be strongly reducing and in the Ni(I) state. From the crystal structure of various forms of the inactive Ni(II) state with the bound coenzymes, it is known that the enzyme has an active site channel which extends from the protein surface deeply into the interior of the protein complex (Ermler et al. 1997). Coenzyme F430 forms the bottom of this channel. Methyl-coenzyme M has to enter this channel before the channel is blocked by coenzyme B. Upon binding, coenzyme B fills the narrowest segment of the channel, with its thiol group facing coenzyme F430 (> Fig. 18.4). The Mcr crystal structure reveals five modified amino acids near the active site. The side chains of specific histidine, arginine, glutamine, and cysteine residues are methylated, and the carbonyl oxygen of a glycine residue is substituted by sulfur. Their high degree of

While the methane formed in the Mcr reaction can be regarded as a waste product, the heterodisulfide product is of central importance for the cell (Hedderich et al. 1998). Reduction of this disulfide is coupled with energy conservation (Deppenmeier et al. 1999; Deppenmeier 2004). Hence, CoM-S-S-CoB can be regarded as the terminal electron acceptor of a respiratory chain in methanoarchaea. The electron donor can be either H2 or coenzyme F420H2, depending on the growth substrate. Evidence that heterodisulfide reduction is coupled to energy conservation comes from studies with Methanosarcina species. In Methanosarcina, all components of the respiratory chain are tightly membrane bound, including a membrane-bound hydrogenase or F420H2 dehydrogenase, the lipophilic electron carrier methanophenazine, and a membrane-bound disulfide reductase (called ‘‘heterodisulfide reductase,’’ Hdr). The latter enzyme functions as a terminal reductase and reduces CoM-S-S-CoB (> Fig. 18.5). Methanophenazine is another novel coenzyme recently discovered in methanogens (Abken et al. 1998). Unlike the other methanogenic coenzymes, methanophenazine seems to be restricted to methanoarchaea belonging to the order Methanosarcinales. The function of this coenzyme is comparable with that of quinones in other respiratory chains.

COOH

O O

H HN H3C H2NOC

5

A 1

N Ni

N D

–

B

N

20

H

CH3

+

O3S

Coenzyme B

Methyl-Coenzyme M

C

15

COO− CH3 H

MCR

N

HOOC

N H

CH3 + HS

COOH 10

H

S

12 13

O COOH –O

O COOH Coenzyme F430

H

COO−

S 3S

N H

S +

Heterodisulfide

CH4

. Fig. 18.3 Structure of coenzyme F430 and the reaction catalyzed by methyl-coenzyme M reductase

CH3 H

OPO3–

OPO3−

Physiology and Biochemistry of the Methane-Producing Archaea

a

18

b CoB-SH

CoM-S-S-CoB CoM-SH

F430

F430

. Fig. 18.4 Active site of methyl-coenzyme M reductase with bound coenzymes. (a) Coenzyme B and coenzyme M are bound in the substrate channel. Note that coenzyme M is neither a substrate nor a product. In this structure, coenzyme M probably mimics the binding position of methyl-coenzyme M with respect to the binding of the sulfonate moiety but not with respect to the binding mode of the thiol group. A Ni-S-CoM species is not considered to be an intermediate in the catalytic cycle. (b) Heterodisulfide (CoM-S-S-CoB) product is bound in the active site channel. The regions indicated with solid lines are the substrate channels near the active site

–

+

CM

VhoA

H2

[NiFe]

Cytoplasm

2 H+ VhoC 3 [4Fe-4S] VhoG Heme b

2 H+

MPH2 CoM-SS-CoB + 2 H+ HS-CoM + HS-CoB

N

Methanophenazinereducing hydrogenase

MP 2 H+

2 [4Fe-4S] 10C HdrD

Heme b HdrE

Heterodisulfide reductase

O

N + 2[H] H N

O

C25H43

N H

. Fig. 18.5 Schematic representation of the respiratory chain catalyzing the reduction of CoM-S-S-CoB by H2 in Methanosarcina species and structure of methanophenazine. Abbreviations: CM, cytoplasmic membrane; [NiFe], active site of the hydrogenases; [4Fe-4S], [4Fe-4S] clusters; heme b, subunits VhoC and HdrE, each containing two heme b binding sites; 10 C, ten highly conserved cysteinyl residues proposed to form one or two iron-sulfur clusters in the active site of Hdr; MP, oxidized methanophenazine; and MPH2, reduced methanophenazine

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Physiology and Biochemistry of the Methane-Producing Archaea

Methanophenazine is reduced by one of the dehydrogenases, a membrane-bound [NiFe] hydrogenase or F420H2 dehydrogenase. For the [NiFe] hydrogenase, two isoenzymes are known. They are both anchored in the membrane via a b-type cytochrome (Deppenmeier et al. 1995, 1999). The catalytic site is most probably facing the extracytoplasmic space, thus releasing two H+ to the outside of the cell after H2 oxidation. This enzyme, which in the past has been called ‘‘methylviologenreducing hydrogenase’’ or ‘‘F420-nonreducing hydrogenase,’’ has now been designated ‘‘methanophenazine-reducing hydrogenase’’ (> Chap. 4, ‘‘H2-Metabolizing Prokaryotes’’ in this volume). The F420H2 dehydrogenase (Fpo) is a multisubunit enzyme composed of six hydrophilic and seven integral membrane proteins (Ba¨umer et al. 2000; > Fig. 18.6). The hydrophilic subunits are facing the cytoplasm. Eleven out of 13 subunits of the enzyme reveal high sequence similarity to subunits of the energy-conserving reduced nicotinamide adenine dinucleotide (NADH):quinone oxidoreductase family (complex I), in particular to the bacterial enzyme which is formed by 14 subunits. Both enzymes differ with respect to the electron input module. In complex I, the electron input module (also called ‘‘NADH-dehydrogenase fragment’’) is formed by three subunits, which oxidize NADH and transfer the electrons to a central module. In Fpo, the NADH-dehydrogenase fragment is replaced by a single subunit, which oxidizes F420H2 (Bru¨ggemann et al. 2000). Reduced methanophenazine generated in the hydrogenase or F420H2 dehydrogenase reaction donates electrons to heterodisulfide reductase (Hdr), which catalyzes the reduction

of CoM-S-S-CoB (Hedderich et al. 1998). In Methanosarcina species, this enzyme is formed by two subunits, a membraneanchoring b-type cytochrome and a hydrophilic catalytic subunit, which is facing the cytoplasm. Unlike most other disulfide reductases, Hdr contains an iron-sulfur cluster in its active site which mediates the reductive cleavage of the disulfide substrate in two one-electron steps (Madadi-Kahkesh et al. 2001; Duin et al. 2003). Using inverted membrane vesicles of Ms. mazei, both the H2: CoM-S-S-CoB oxidoreductase and the F420H2:CoM-S-S-CoB oxidoreductase reactions were found to be coupled with the transfer of 4H+/2e across the cytoplasmic membrane. More recent studies revealed that each partial reaction, the reduction of methanophenazine by H2 or F420H2 and the reduction of CoM-S-S-CoB by reduced methanophenazine, is coupled to the translocation of 2H+/2e (Ide et al. 1999; Ba¨umer et al. 2000). As shown in > Fig. 18.5, H+-translocation in both the hydrogenase- and the heterodisulfide reductase reactions could function via a redox-loop mechanism. This mechanism cannot apply for the F420H2 dehydrogenase. Like complex I, this enzyme is thought to function as a proton pump (Ba¨umer et al. 2000). In the hydrogenotrophic methanoarchaea belonging to the four other orders of methanoarchaea, the respiratory chain catalyzing the reduction of CoM-S-S-CoB differs significantly from that described above for Methanosarcina. These organisms do not contain heme. Thus, b-type cytochromes can be excluded as membrane anchors and electron carriers of membranebound dehydrogenases and reductases. Furthermore, methanophenazine has not been detected in these organisms (U. Deppenmeier, personal communication). As deduced from

NADH + H+ F F420H2

F FAD NAD+

N

D

C

4Fe

4Fe I

I 2 H+

Nuo O 2Fe

J

2Fe 4Fe

4Fe

D

2 H+

K

2Fe E

C

4Fe

4Fe 4Fe

Fpo

4Fe

4Fe

F420

G

FMN

M

4Fe

L MPH2

Q

B

MP

H

A

K

J

N

4Fe

L

M

B

H

A

QH2

. Fig. 18.6 Comparison of the structures of F420H2 dehydrogenase (Fpo) from Methanosarcina mazei and NADH:quinone oxidoreductase (Nuo) from Escherichia coli. Capital letters indicate subunits of the enzymes. Abbreviations: 4Fe, [4Fe-4S] cluster; 2Fe, [2Fe-2S] cluster; FMN, flavin mononucleotide, FAD, flavin dinucleotide; Q, ubiquinone or menaquinone; MP, oxidized methanophenazine; MPH2, reduced methanophenazine. FpoO has no counterpart in complex I, and its function is not known

Physiology and Biochemistry of the Methane-Producing Archaea

genome sequences, the membrane-bound F420H2dehydrogenase (Fpo) is also lacking from these organisms. Reduction of CoM-S-S-CoB in a non-Methanosarcina species has mainly been studied in Methanothermobacter marburgensis, which belongs to the order Methanobacteriales. In this organism, heterodisulfide reductase, which is composed of three hydrophilic subunits, forms a tight and catalytically active complex with a [NiFe] hydrogenase (Setzke et al. 1994; Hedderich et al. 1998). This complex was designated as ‘‘H2: heterodisulfide oxidoreductase complex.’’ Hdr from this organism shares the catalytic subunit with the enzyme from Methanosarcina but is lacking a membrane anchor (Hedderich et al. 1998). The hydrogenase present in this complex is also lacking a membrane subunit and therefore is clearly different from the methanophenazine-reducing hydrogenase present in Methanosarcina species. Hence, the six subunits of the H2: heterodisulfide oxidoreductase complex are all hydrophilic. The three transcriptional units encoding the different subunits of the complex do not contain additional open reading frames (ORFs) encoding potential integral membrane proteins, which might have been separated from the hydrophilic part during the purification. Hence, there is at present no conclusive answer how this apparently cytoplasmic protein complex can couple the reduction of CoM-S-S-CoB by H2 with the generation of a proton motive force.

2002). The enzyme catalyzes the reversible dehydrogenation of formylmethanofuran to CO2 and methanofuran. In vitro, this enzyme can be assayed with viologen dyes as artificial electron donors or acceptors. Fmd purified from Methanosarcina barkeri is a membrane-associated molybdenum iron-sulfur protein composed of six subunits. Other organisms, such as Methanothermobacter species, contain two Fmds, one enzyme containing molybdenum and the second enzyme containing tungsten bound to the molybdopterin cofactor. The electron donor to Fmd and the hydrogenase participating in this reaction were unknown until recently. Upon identification of a novel membrane-bound [NiFe] hydrogenase in Methanosarcina barkeri, called the ‘‘energy-converting hydrogenase’’ or Ech, it was suggested that this enzyme could play an essential role in driving this endergonic redox reaction (Ka¨nkel et al. 1998; Meuer et al. 1999). Ech is a multisubunit membranebound [NiFe] hydrogenase. The six subunits of the enzyme exhibit high sequence similarity to subunits of the energyconserving NADH:quinone oxidoreductase (complex I) of mitochondria and bacteria (Hedderich 2004). In vitro, Ech catalyzes the reversible reduction of a special ferredoxin containing two [4Fe-4S] clusters by H2 (Meuer et al. 1999). The oxidation of formylmethanofuran to CO2 by the M. barkeri membrane fraction is also ferredoxin dependent, and ferredoxin was therefore proposed to function as an electron carrier between Ech and Fmd in vivo. Support for this hypothesis came from the physiological characterization of a Dech mutant. This mutant was unable to make Ech and unable to form formylmethanofuran and reduce CO2 to methane (Meuer et al. 2002). In addition, the Dech mutant was unable to biosynthesize acetyl-CoA and pyruvate via the acetyl-CoA synthase and pyruvate oxidoreductase reactions, respectively, using H2 as the electron donor. Like the reaction catalyzed by Fmd, these reactions require a strong reductant. These data support the model depicted in > Fig. 18.8. According to this scheme, Ech catalyzes the reduction of a low-potential ferredoxin by H2. Reduced ferredoxin can then function as the electron donor of Fmd but also of other oxidoreductases which require a low-potential electron donor. Since Ech is tightly membrane bound via two integral membrane subunits and resembles the central part of complex I, it was suggested that ferredoxin reduction is driven by reversed

Reductive Activation of CO2 to Formylmethanofuran The reduction of CO2 to formylmethanofuran (CHO-MFR) is the first step of the methanogenic pathway from H2/CO2 (> Fig. 18.7). This reaction is highly endergonic with H2 as electron donor (DG0 = +16 kJ·mol 1). It becomes even more endergonic (DG0 = +45 kJ·mol 1) under the low hydrogen partial pressures prevailing in the natural habitats of methanoarchaea. In cell suspension experiments, it was shown that this reduction is driven by reversed electron transport and requires an electrochemical ion gradient (Kaesler and Scho¨nheit 1989; Deppenmeier et al. 1996). A key enzyme involved in the catalysis of the reaction (> Fig. 18.7) is formylmethanofuran dehydrogenase (Fmd; for a review, see Vorholt and Thauer

R

R

H

+

O

N

NH3 + H2 + CO2

O

COO–

C

–OOC

COO

O

N

–

H

. Fig. 18.7 Reaction catalyzed by formylmethanofuran dehydrogenase

C N O

O C

+ H2O + H+

H

O

H

COO–

18

C COO–

N H

O CH2

R

645

646

18

Physiology and Biochemistry of the Methane-Producing Archaea

– CO2+ MFR

H4MPT formyltransferase (Ftr), forming N5-formyl-H4MPT (reaction 1; Shima et al. 2002):

cytoplasmicmembrane +

Formyl MFR þ H4 MPT $ MFR þ formyl H4 MPT ð18:1Þ

Ech 2 Fdred 4Fe F 4Fe

Fmd

B C 4Fe

ΔµH + ? A

2 Fdox

ΔµNa + ?

E [NiFe]

Formyl H4 MPT þ H D

CHO–MFR

2H+

This is different from folate biochemistry where only N10-formyl-H4F is known. A cyclohydrolase then converts N5-formyl-H4MPT to N5,N10-methenyl-H4MPT (reaction 2; Shima et al. 2002):

H2

. Fig. 18.8 Proposed function of Ech hydrogenase in the first step of methanogenesis from H2/CO2 in Methanosarcina barkeri. Whether Ech hydrogenase uses a H+- or Na+-motive force to drive the reduction of the ferredoxin by H2 remains to be demonstrated. Abbreviations: Fmd, formylmethanofuran dehydrogenase; Fd, 2 [4Fe-4S] ferredoxin; MFR, methanofuran; CHO-MFR, formylmethanofuran

electron transport. In contrast, the ferredoxin-dependent reduction of CO2 to formylmethanofuran is not directly linked to an electrochemical ion gradient. This conclusion is supported by further experiments with the Dech mutant. In this mutant, CH4 formation from H2/CO2 can be restored by CO or pyruvate, two strong electron donors which can couple to Fmd (Stojanowic and Hedderich 2004). Moreover, when H2 was replaced by CO as electron donor for the first step of methanogenesis, CH4 formation was no longer dependent on an energized membrane. Thus, reduction of the electron carrier by H2, which is catalyzed by Ech, and not the Fmd reaction per se, is dependent on an energized membrane.

Tetrahydromethanopterin Replaces Tetrahydrofolate in the C1 Pathway of Methanoarchaea Tetrahydromethanopterin (H4MPT) carries the one-carbon unit at the oxidation level of formate, formaldehyde, and methanol. In Methanococcales, Methanosarcinales, and Methanomicrobiales, derivatives of H4MPT such as tetrahydrosarcinapterin (H4SPT) are used instead of H4MPT (DiMarco et al. 1990; Gorris and van der Drift 1994). H4MPT is an analogue of tetrahydrofolate (H4F), which is the C1 carrier used by most other organisms (> Fig. 18.9). The most important structural differences between the coenzymes are that H4F has an electronwithdrawing carbonyl group conjugated to the N10 via the aromatic ring and H4MPT has methyl groups at the ring carbons C7 and C11 (Maden 2000). The chapter into the H4MPTdependent reaction cascade is catalyzed by formyl-MFR:

$ methenyl H4 MPT þ H2 O ð18:2Þ

The following reduction to methylene-H4MPT is catalyzed by N5,N10-methylene-H4MPT-dehydrogenase using reduced coenzyme F420 (F420H2) as electron donor (reaction 3; Hagemeier et al. 2003): Methenyl H4 MPT þ F420 H2 $ methylene H4 MPT þ F420 þ H ð18:3Þ 5

10

This reaction is analogous to the N , N -methyleneH4F-dehydrogenase reaction, although the electron donor is different. As will be described below, some methanoarchaea have an alternative enzyme which catalyzes methenyl-H4MPT reduction with H2 as electron donor. N5,N10-methylene-H4MPT is then reduced to N5-methyl-H4MPT by N5,N10-methyleneH4MPT reductase using F420H2 as electron donor (reaction 4; Shima et al. 2002): Methylene H4 MPT þ F420 H2 $ methyl H4 MPT þ F420 ð18:4Þ In contrast to the corresponding enzyme of the H4F-dependent pathway, the reductase from methanoarchaea lacks a flavin. In the H4MPT pathway, the methylene-H4MPT and methenyl-H4MPT redox couples are substantially more negative than in the H4F pathway (> Fig. 18.9). This enables reversible coupling to the low redox-potential cofactor F420 (> Fig. 18.10). This could be one reason why methanoarchaea utilize methanopterin instead of tetrahydrofolate in their C1 pathway (Maden 2000). F420 is a 5-deazaflavin, which is responsible for the blue-green fluorescence of methanoarchaea because of its high abundance in the cell. Unlike flavins, F420 only catalyzes hydride transfer reactions. Thus, it functions as a two-electron donor like NAD(P)+. The E0 of the F420/F420H2 couple is, however, about 360 mV, compared with 320 mV for the NAD(P)+/NAD(P)H couple (Warkentin et al. 2001).

A Sodium Ion Pumping Methyltransferase The last H4MPT-dependent reaction in the methanogenic C1 pathway is the transfer of the methyl group from N5-methylH4MPT to coenzyme M (> Fig. 18.11). This exergonic reaction (DG0 = 30 kJ·mol 1) is catalyzed by N5-methyl-H4MPT: CoM-SH methyltransferase (Mtr), a membrane-integral

Physiology and Biochemistry of the Methane-Producing Archaea

H O

5

N

H

COO–

H

N

N

H2N

10

N

N

COO–

N

H

H

O

18

H4F

H

HO HO

H O H

O

COO–

P O O

O

C N H

COO–

H4MPT

H

ΔE°‘ H

ΔE°‘

– 360 mV C

OH

H CH3 H

N

COO–

O–

O OH

H CH3

5

N

OH

10

N

N H 2N

O N

H

N

H

H C

– 300 mV

N +

N

– 330 mV

H

– 200 mV

CH3 N

N

N

Methylene-H4MPT Methylene-H4F

Methenyl-H4MPT Methenyl-H4F

Methyl-H4MPT Methyl-H4F

. Fig. 18.9 Structure of tetrahydromethanopterin (H4MPT), tetrahydrofolate (H4F), and their C1-derivatives. Methanosarcina species contain a H4MPT analogue called ‘‘tetrahydrosarcinapterin’’ (H4SPT) in which the a carboxyl group of the side chain is linked to a glutamate residue (highlighted in magenta)

O H

H

O H

N

O

N

N

OH

O

F420

O

R= OH

N

N

H

R

OH

F420H2 O

OH

H

N

R

OH

H

O

COO–

O

H N

O

P –

CH3

N H

O

COO– COO–

. Fig. 18.10 Structure of coenzyme F420 in the oxidized and the reduced state. The structural similarity to pyridine nucleotides is indicated. Methanothermobacter thermautotrophicus and Methanococcus voltae F420 contain two glutamyl residues (F420-2), and Methanosarcina barkeri contains F420 with four and five glutamyl residues (F420-4,5)

multienzyme complex composed of eight different subunits (MtrA-H; for a review, see Gottschalk and Thauer 2001; > Fig. 18.11). The enzyme is strictly dependent on sodium ions (Weiss et al. 1994). Subunit MtrA harbors a cob(I)amide prosthetic group (Ga¨rtner et al. 1993), which is methylated and demethylated during the catalytic cycle. The demethylation reaction is Na+-dependent. After reconstitution into proteoliposomes, the enzyme was shown to pump Na+ (Lienard et al. 1996). A ratio of 1.7 mol of Na+ translocated per mol of methyl group transferred was determined. Thus, the enzyme appears to be a sodium pump that couples the methyl transfer to coenzyme M with formation of a sodium motive force. The bound cobamide may play an important role in the enzyme mechanism. In aqueous solution, cob(II)amide and methylcob(III)amide contain axial ligands to the cobalt, whereas cob(I)amide does not (Kra¨utler 1998). In unmethylated MtrA, the bound cob(I)amide should therefore have no axial ligand. Upon methylation of cob(I)amide to methylcob(III)amide, the methylcob(III)amide is expected to bind a histidine residue of

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Physiology and Biochemistry of the Methane-Producing Archaea

Na+

MtrE

CM

MtrC

Cytoplasma

MtrB

MtrD

D

MtrG

MtrA MtrF

S Co

CH3-H4MPT

O

Zn2+ S

R

N

SO3–

H3C

N

HN H2N

CH3

H N

SO3– H-S-CoM

H

CH3-S-CoM

N

MtrH

H O

H4MPT

H N

R

N

HN H2N

H

N

N H

. Fig. 18.11 Model of the methyl-H4MPT:CoM-SH methyltransferase complex. A conserved aspartate residue (D) predicted to be located in a transmembrane helix of subunit MtrE is highlighted. This residue could be essential for sodium ion translocation (Modified from Gottschalk and Thauer (2001))

the protein as an axial ligand (Harms and Thauer 1997; > Fig. 18.12). Binding of this axial ligand could therefore be associated with a conformational change of the protein. Upon demethylation of the cobamide, the axial ligand would be lost, and the conformational change would be reversed. Since demethylation is Na+ dependent, the conformational change associated with this step can be coupled with the vectorial translocation of Na+. The MtrH subunit can be separated from the MtrA-H-complex. The isolated MtrH subunit can catalyze the methylation of free cob(I)amide with methyl-H4MPT (Hippler and Thauer 1999). Therefore, MtrH is thought to catalyze the methylation of the corrinoid prosthetic group, which is bound to MtrA. Subunit MtrE is thought to transfer the methyl group from the corrinoid prosthetic group of MtrA to coenzyme M, which is the Na+-dependent reaction (Gottschalk and Thauer 2001). MtrE is predicted to form six transmembrane spanning helixes and to have a large cytoplasmic domain containing a typical zinc-binding motif. All enzymes known to date that catalyze the alkylation of a thiol group are zinc proteins. The reaction catalyzed by N5-methyl-H4MPT:CoMSH methyltransferase is analogous to the formation of

Unmethylated MtrA

Methylated MtrA CH3-H4MPT

H4MPT CH3

MtrH

Co3+

MtrE ?

His

Co1+

His CH3-s-CoM

H-s-CoM Na+

. Fig. 18.12 Proposed conformational change of subunit MtrA of the methylH4MPT:CoM-SH methyltransferase complex upon methylation and demethylation of its corrinoid prosthetic group. The MtrH subunit is proposed to catalyze the methylation of the cobamide bound to MtrA. The demethylation reaction is thought to be catalyzed by MtrE and coupled with vectorial sodium ion translocation since this reaction is sodium ion dependent (Modified from Gottschalk and Thauer (2001))

Physiology and Biochemistry of the Methane-Producing Archaea

methionine from N5-methyl-H4F and homocysteine, which is catalyzed by methionine synthase (Banerjee et al. 1989). However, methionine synthase is a soluble enzyme containing only one type of subunit, reflecting the fact that the methyl transfer to homocysteine is not coupled to energy conservation.

Activation of Methanol and Methylamines As shown in > Fig. 18.13, methylotrophic methanogenesis begins with the transfer of the methyl group from a variety of substrates to coenzyme M. For each substrate, there is a different methyltransferase system, specific for methanol (Mta), monomethylamine (Mtm), dimethylamine (Mtb), trimethylamine (Mtt), tetramethylammonium (Mtq), and methylthiols (Mts; Thauer and Sauer 1999; Ferguson et al. 2000). Each system is composed of two methyltransferases, designated ‘‘MT1’’ (MtaB, MtmB, MtbB, MttB, and MtqB) and ‘‘MT2’’ (MtaA, MtbA, and MtqA), and a substrate-specific methylotrophic corrinoid protein (MtaC, MtmC, MtbC, MttC, and MtqC) containing a modified cobamide. MT1 in each system catalyzes the methylation of the reduced corrinoid protein, and MT2 catalyzes the transfer of the methyl group from the corrinoid protein to coenzyme M. Only in dimethylsulfide:coenzyme M methyltransferase are both methyl transfer reactions catalyzed by the same subunit (MtsA; Tallant et al. 2001). The MT2 proteins have high sequence similarity and contain zinc in the active site. Likewise, the sequences of the corrinoid proteins are related, all exhibiting a corrinoid-binding

motif. In contrast, the substrate-activating MT1 enzymes are not phylogenetically related. For instance, MtaB, which activates methanol, is a zinc protein, but the other methylamine methyltransferases are not (Sauer and Thauer 1997). The genes encoding MtmB, MtbB, and MttB contain a single conserved in-frame amber codon (UAG) that is read through during translation (James et al. 2001). In the structure of MtmB, the UAG-encoded residue was identified as a lysine in amide linkage to (4R, 5R)-4-substituted-pyrroline-5-carboxylate (called ‘‘pyrrolysine’’; Hao et al. 2002). Furthermore, an amber-decoding tRNA was identified (Srinivasan et al. 2002). Pyrrolysine can therefore be regarded as the twenty-second genetically encoded amino acid. Pyrrolysine is thought to position the methyl group of methylamine for attack by the corrinoid protein (Hao et al. 2002).

The Aceticlastic Reaction Species of Methanosarcina, as well as those of Methanosaeta, grow during the catabolism of acetate to CO2 and CH4 (Ferry 1997). This is the acetate cleavage or aceticlastic reaction, where methane is formed without oxidation of the methyl group of acetate. Instead, after activation to acetyl-CoA, the acetyl C-C bond is cleaved by the multienzyme complex of acetyl-CoA synthase and carbon monoxide dehydrogenase (Acs/CODH), which in Methanosarcina barkeri and Methanosarcina thermophila is composed of five different subunits (a subunit, CdhA; b subunit, CdhC; g subunit, CdhE; d subunit, CdhD; and subunit, CdhB). The overall reaction catalyzed by the

CH3-S-CoM

H-S-CoM

H2O Methanol

MMS

Co

MtsA

MtaA

MtaC

DMS

Co

MtaB

MtsB

NH4+

MtbA

MtqA

Co

MtqC

MtmC

MMA

18

MtbA

MtaA/ MtbA

MtmB MMA

MttC Co

Co MtbC

Co TMA

MtqB

DMA QMA DMA

MttB

MtbB TMA

. Fig. 18.13 Enzymes involved in the formation of methyl-coenzyme M from methanol, monomethylamine (MMA), dimethylamine (DMA), trimethylamine (TMA), tetramethylammonium (QMA), and dimethylsulfide (DMS). Except for DMS, the B subunits transfer the methyl groups from the substrates to the corrinoid prosthetic groups of the C subunits. The A subunits then transfer the methyl groups from the corrinoid to CoM-SH. For DMS, the A subunit catalyzes both transfers. Abbreviations: MMS, methanethiol; Co, corrinoid prosthetic group

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Physiology and Biochemistry of the Methane-Producing Archaea

complex is the conversion of acetyl-CoA and tetrahydrosarcinapterin (H4SPT) to CO2, N5-methyl-tetrahydrosarcinapterin (CH3-H4SPT), CoA-SH, and reducing equivalents (reaction 5). Tetrahydrosarcinapterin is similar in structure and function to tetrahydromethanopterin, which is common in the hydrogenotrophic methanogens. Acetyl

CoA þ H4 SPT þ H2 O þ 2Fdox $ CoA þ CH3 H4 SPT þ CO2 þ 2FDred þ 2H

CH3COO− ATP CH3CO~SCoA H4SPT

+

−

Fdox

SH

þ

Acs/CODH

ð18:5Þ A ferredoxin was identified as the physiological electron acceptor. In autotrophic methanoarchaea and the homoacetogenic bacteria like Moorella thermoacetica, a homologous enzyme system functions in the reverse direction for the biosynthesis of acetyl-CoA. This overall reaction is made up of a series of partial reactions catalyzed by different protein subcomponents of the complex (Abbanat and Ferry 1991; Grahame and DeMoll 1996). The b subunit, the recombinant form of which can be produced in Escherichia coli, reacts with acetyl-CoA to form an acetylenzyme intermediate. Furthermore, this subunit catalyzes the formation of acetyl-CoA from CoA-SH, CO, and methylcobalamin in the absence of other Acs/CODH subunits, demonstrating that this subunit catalyzes the reversible C–C bond activation (Gencic and Grahame 2003). The b subunit also harbors the ‘‘A-cluster,’’ which contains a Ni-Ni-[4Fe-4S] site, as deduced from the crystal structures of Acs/CODH from Moorella thermoacetica (Darnault et al. 2003; Seravalli et al. 2004) and Carboxydothermus hydrogenoformans (Svetlitchnyi et al. 2004). The CO generated in the C–C cleavage reaction is transferred via a gas channel to the site of the CO dehydrogenase activity, which is on the a subcomplex. The isolated a subcomplex catalyzes the oxidation of CO to CO2. Furthermore, the sequence of the a subunit is related to the sequences of the much simpler CO dehydrogenases from Rhodospirillum rubrum and Carboxydothermus hydrogenoformans. The active site of CO dehydrogenase also contains a Ni-Fe/S center, which could be either a [Ni-Fe4-S4] or a [Ni-Fe4-S5] center, as deduced from the crystal structure of these enzymes (Dobbek et al. 2001; Drennan et al. 2001). The methyl group generated in the b subunit is transferred to the corrinoid cofactor present in the gd subcomplex, which catalyzes the subsequent methyl transfer to the substrate H4SPT. Here, the methyl group enters the general methanogenic pathway, which leads to the formation of CH4 (> Fig. 18.2). Reducing equivalents required for the reduction of the heterodisulfide are provided by reduced ferredoxin formed in the CO dehydrogenase reaction. There might be alternative electron transport chains to couple ferredoxin oxidation to heterodisulfide reduction. In Methanosarcina barkeri, H2 is thought to be an intermediate in this electron transfer reaction. This conclusion is based on several observations. First, H2 accumulates during growth on acetate. Second, acetate-grown cells have high levels of the Ech

CO2 + CoA

ΔµH+ ?

Ech

Fdred 2H+

H2

H2

CH3-H4SPT Vho CH3-CoM

H-S-CoM

MPH2

2 H+

MP

H-S-CoB Hdr CoM-S-S-CoB +2H+ CH4

. Fig. 18.14 Pathway of methanogenesis from acetate in Methanosarcina barkeri. Recent data indicate that the 2 [4Fe-4S] ferredoxin (Fd) from M. barkeri mediates electron transfer between acetyl-CoA synthase/CO dehydrogenase (Acs/CODH) and Ech hydrogenase. Abbreviations: CH3-H4SPT, methyl-tetrahydrosarcinapterin; MP, methanophenazine; MPH2, reduced methanophenazine

hydrogenase and methanophenazine-reducing hydrogenase. Third, Ech hydrogenase is essential for growth of M. barkeri on acetate. It has therefore been proposed that this enzyme catalyzes H2 formation from reduced ferredoxin (Meuer et al. 2002). H2 thus formed could then diffuse to the extracytoplasmic side of the membrane, where it becomes oxidized by the methanophenazine-reducing hydrogenase. Reduced methanophenazine is then the electron donor for the heterodisulfide reductase (> Fig. 18.14). On the other hand, M. acetivorans forms methane from acetate but lacks a functional Ech hydrogenase (Galagan et al. 2002). Hence, there must exist an alternative route to channel electrons from reduced ferredoxin into a membranebound electron transport chain that leads to heterodisulfide reduction.

The Hydrogenases of Methanoarchaea: A Summary For most methanoarchaea, methanogenesis from H2 and CO2 is the only way to obtain energy for growth. Also growth on acetate could involve H2 formation and H2 consumption as discussed above. Therefore, hydrogenases are essential enzymes for methanoarchaea, which is reflected by the presence of

Physiology and Biochemistry of the Methane-Producing Archaea

five different types of hydrogenases in these organisms. Four of these enzymes are [NiFe] hydrogenases, and one enzyme is an iron-sulfur cluster-free hydrogenase that has only been found in methanoarchaea. Methanoarchaea seem to be lacking [FeFe] hydrogenases. For a more detailed description of hydrogenases including those from methanoarchaea, see > Chap. 4, ‘‘H2-Metabolizing Prokaryotes’’ in this volume.

F420-Reducing Hydrogenase This enzyme (Frh) is conserved in all methanoarchaea studied. Some organisms contain two closely related isoenzymes. The enzyme catalyzes the reduction of the deazaflavin coenzyme F420 and thus provides the reducing equivalents for the two intermediate reduction steps of the C1 pathway. Frh is a soluble [NiFe] hydrogenase composed of three subunits, including the ‘‘hydrogenase large subunit’’ and the ‘‘hydrogenase small subunit’’ that form the basic module of all [NiFe] hydrogenases. The third subunit contains iron-sulfur clusters and FAD. It is assumed to harbor the F420-binding site (Sorgenfrei et al. 1997).

H2-Forming Methylene-H4MPT Dehydrogenase As outlined above, all hydrogenotrophic methanogens possess an F420-dependent dehydrogenase for the reduction of methenyl-H4MPT. Reduction of F420 to F420H2 by H2 is catalyzed by Frh. Methanoarchaea belonging to the orders Methanobacteriales, Methanococcales, and Methanopyrales also possess an enzyme that directly reduces methenyl-H4MPT to methylene-H4MPT using H2 as the electron donor (Thauer et al. 1996; > Fig. 18.15). Because this enzyme oxidizes H2, it is a hydrogenase by definition. However, because this reaction is so unusual, it has been called the ‘‘H2-forming methylene-H4MPT dehydrogenase’’ (Hmd). In contrast to the well-characterized [NiFe] hydrogenases and [FeFe] hydrogenases, Hmd does not contain Ni or iron-sulfur clusters. The primary sequence of Hmd does not possess similarity to known proteins.

O

+ N H2N

N

pro-R pro-S H O N

Hmd H

N

Furthermore, the enzyme is not inhibited by CO at concentrations known to inhibit other hydrogenases, and it does not catalyze the reduction of redox dyes such as benzyl- or methylviologen. It does catalyze the exchange between H2 and protons and the conversion of para H2 to ortho H2 but only in the presence of methenyl-H4MPT. More detailed mechanistic studies have shown that the enzyme catalyzes the reversible reduction of methenyl-H4MPT to methylene-H4MPT in a ternary complex catalytic mechanism. In this reaction, a hydride is transferred from H2 into the pro-R position at C14 of methenyl-H4MPT. Iron at concentrations up to 1 mol of Fe per mol of enzyme is the only metal that has been detected in Hmd. This iron was not redox active and not considered to be functional. The enzyme was therefore called ‘‘metal-free’’ hydrogenase. Recently, active enzyme was shown to contain a cofactor (Buurman et al. 2000). Addition of the purified cofactor to the apoprotein, which can be produced in E. coli, resulted in active enzyme. The structure of the active cofactor is not yet known. But upon illumination with ultraviolet (UV)-A/blue light, the cofactor is inactivated and Fe and CO are released (Lyon et al. 2004b). The remaining organic component could be cleaved by phosphodiesterase to GMP and a pyridone moiety, which is a new structure in biology (Shima et al. 2004). How this organic compound is involved in iron complexation in the active Hmd cofactor remains to be shown. There is experimental evidence that two CO are bound to the iron center (Lyon et al. 2004a). Interestingly, CO is also a ligand to the iron center in [NiFe] and [FeFe] hydrogenases. In cells cultivated under Ni-limiting conditions, the [NiFe] hydrogenase Frh is barely detectable, while the concentration of Hmd in the cell increases (Afting et al. 1998). Hmd in combination with F420-dependent methylene-H4MPT dehydrogenase (Mtd) mediates the reduction of coenzyme F420 by H2 and thus provides an alternative source for reduced coenzyme F420. This allows the cell to spare Ni. In contrast to the [NiFe] hydrogenase, Frh, Hmd, and Mtd are not oxygen sensitive. This becomes important in the context of the recent finding that methanoarchaea contain an F420H2 oxidase, which catalyzes the reduction of O2 to H2O with F420H2 as the electron donor (Seedorf et al. 2004). The reduction of O2 with H2 in

H N N H

CH3 + H2

CH3

Methenyl-H4MPT

18

H2N

CH3+ H+

N

N

N N

N H

CH3

Methylene-H4MPT

. Fig. 18.15 Reaction catalyzed by H2-forming methylene-H4MPT dehydrogenase (Hmd). In the presence of H2, methenyl-H4MPT is reduced to methylene-H4MPT

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Physiology and Biochemistry of the Methane-Producing Archaea

methanoarchaea is not coupled with energy conservation. The function of this oxidase is most probably to reduce the intracellular O2 concentration to a level that allows growth and methanogenesis. There is evidence that the O2 concentration has to be lowered well below 5 mM in order for a ‘‘nanaerobe’’ to grow (Baughn and Malamy 2004). The function of F420H2 oxidase is, therefore, O2 detoxification.

F420-Nonreducing Hydrogenase F420-nonreducing hydrogenase (Mvh) is a soluble [NiFe] hydrogenase. In addition to the basic hydrogenase module of two subunits, the enzyme contains a third subunit, a 17-kDa protein that carries a [2Fe-2S] cluster. In M. marburgensis, Mvh forms an enzyme complex with heterodisulfide reductase (Hdr). There is indirect evidence that the hydrogenase interacts via its 17-kDa subunit with Hdr (Stojanowic et al. 2003). This type of hydrogenase is not found in Methanosarcina species.

Methanophenazine-Reducing Hydrogenases Methanosarcina species form two closely related [NiFe] hydrogenases, encoded by the vho and the vht transcriptional units. In addition to the basic hydrogenase module, these enzymes contain a membrane-anchoring b-type cytochrome, which easily becomes separated from the hydrogenase module during purification. These enzymes possess the highest similarity to the membrane-bound, periplasmically oriented uptake hydrogenases of bacteria (Vignais et al. 2001). Vho and vht also contain a twin arginine leader peptide in their hydrogenase small subunit, indicating that the hydrophilic subunits of these enzymes are translocated across the membrane by twin arginine translocation (TAT) machinery. This type of hydrogenase has only been found in Methanosarcina species where it is part of the H2:CoM-S-S-CoB oxidoreductase system (Deppenmeier et al. 1999; > Fig. 18.5). The vhoGAC operon is expressed during growth on H2/CO2, methanol, or acetate. The vhtGAC operon is only expressed during growth on H2/CO2 and methanol but not during growth on acetate (Deppenmeier 1995). Whether this pattern of expression reflects a different metabolic function is not known.

Energy-Converting [NiFe] Hydrogenases Energy-converting [NiFe] hydrogenase (Ech) is an integral membrane protein, which, when purified, is composed of six subunits, corresponding to the products of the echABCDEF operon (Ka¨nkel et al. 1998; Meuer et al. 1999). Ech hydrogenase is only distantly related to the other [NiFe] hydrogenases found in methanoarchaea. The subunits of this enzyme are closely related to members of a small group of membrane-bound [NiFe] hydrogenases, such as hydrogenase 3 from E. coli and the CO-induced hydrogenase from Rhodospirillum rubrum.

The sequences of the six subunits conserved in these enzymes are closely related to subunits present in the central part of complex I from mitochondria and bacteria (Hedderich 2004). The EchA and EchB subunits of the enzyme are predicted to be membrane-spanning proteins, while the other four subunits are expected to extrude into the cytoplasm. A low-potential, soluble two [4Fe-4S] ferredoxin (E00 = 420 mV) isolated from M. barkeri was identified as the electron donor/acceptor of Ech. As outlined above, this enzyme provides the cell with reduced ferredoxin required for the first step of methanogenesis and for certain anabolic reactions. In vivo, the reduction of the ferredoxin by H2 is thought to be driven by reversed electron transport. In aceticlastic methanogenesis, Ech was proposed to catalyze the reverse reaction (i.e., the production of H2 with reduced ferredoxin as electron donor). This has been concluded from experiments with intact cells. Cell suspensions of wild-type M. barkeri convert CO quantitatively to CO2 and H2. Cell suspensions of the Dech mutant catalyzed the oxidative half of the aceticlastic pathway (conversion of CO to CO2 and H2) at a significantly lower rate than the wild type, indicating that Ech is the hydrogenase involved in this reaction (Meuer et al. 2002). Importantly, the conversion of CO to CO2 and H2 in wild-type M. barkeri was found to be coupled to the generation of a proton motive force. This is consistent with the putative ion-translocating activity of Ech. Ech hydrogenase thus far has only been purified from Methanosarcina species. The genomes of Methanothermobacter thermautotrophicus, Methanococcus jannaschii, and Methanopyrus kandleri do not encode a homologue of the sixsubunit Ech hydrogenase present in Methanosarcina. However, these organisms encode related enzymes, which are predicted to have a much more complex subunit architecture (> Fig. 18.16). Methanothermobacter thermautotrophicus, M. marburgensis, and M. jannaschii each encode two hydrogenases of this type, designated ‘‘Eha’’ and ‘‘Ehb’’ (Tersteegen and Hedderich 1999). Methanopyrus kandleri only encodes for one of these hydrogenases (Slesarev et al. 2002). In M. marburgensis, the length of the transcription units was determined. The eha operon (12.5 kb) and the ehb operon (9.6 kb) were found to be composed of 20 and 17 ORFs, respectively. Sequence analysis of the deduced proteins indicated that the eha and ehb operons each encode a [NiFe] hydrogenase large subunit, a [NiFe] hydrogenase small subunit, and two conserved integral membrane proteins. These proteins show high sequence similarity to subunits of Ech hydrogenase from Methanosarcina barkeri. In addition to these four subunits, the eha operon encodes a 6[4Fe-4S] polyferredoxin, a 10[4F-4S] polyferredoxin, four nonconserved hydrophilic subunits, and ten nonconserved integral membrane proteins; the ehb operon encodes a 2[4Fe-4S] ferredoxin, a 14[4Fe-4S] polyferredoxin, two nonconserved hydrophilic subunits, and nine nonconserved integral membrane proteins. Since Methanothermobacter species only grow with H2/CO2 as energy substrates, it has been proposed that these membrane-bound [NiFe] hydrogenases catalyze the reduction of a low-potential ferredoxin or polyferredoxins by H2 in a reaction driven by

Physiology and Biochemistry of the Methane-Producing Archaea

Ech hydrogenase

ech

A

B

C D

E

F

H I

J

18

M. barkeri Eha hydrogenase

eha A B C D E F

G

KL M N

O

P

Q

R

S

T

M. marburgensis

Ehb hydrogenase

ehb A B C D E

F

G H I J

K

L M

N

O

P Q

M. marburgensis

= hydrogenase large subunit = hydrogenase small subunit; 1 × [4Fe-4S] = electron-transfer protein; n × [4Fe-4S]

= integral membrane protein ( = conserved integral membrane protein) = non-conserved hydrophilic protein

. Fig. 18.16 Organization of the Methanosarcina barkeri ech operon and the Methanothermobacter marburgensis eha and ehb operons. Abbreviations: [4Fe-4S], iron-sulfur cluster; n x [4Fe-4S], polyferredoxin encoded by the operon

reversed electron transport, in analogy to the function of Ech hydrogenase in M. barkeri when the organism is cultivated on H2/CO2. A purification of these enzymes has not been achieved thus far.

Methanogenic Coenzymes and Enzymes in Nonmethanogenic Archaea and Bacteria Sulfate-Reducing Archaea Use Three Methanogenic Coenzymes for the Oxidation of Reduced C1 Compounds to CO2 So far, all isolated archaeal sulfate reducers belong to the genus Archaeoglobus. The best-studied species is Archaeoglobus fulgidus, for which the genome sequence is also known (Klenk et al. 1997). Archaeoglobus fulgidus couples the oxidation of lactate to CO2 with the reduction of sulfate to H2S. Lactate is first oxidized to pyruvate, which is subsequently converted to acetyl-CoA, CO2, and 2[H]. Cleavage of the C–C-bond of acetyl-CoA is catalyzed by the Acs/CODH complex, which has the same subunit architecture and high sequence similarity to the enzyme from methanoarchaea (Dai et al. 1998). This reaction generates enzyme-bound CO, which is oxidized to CO2, and an enzyme-bound methyl group. For the oxidation of the methyl group to CO2, A. fulgidus uses three coenzymes characteristic of the methanoarchaea: tetrahydromethanopterin, methanofuran, and coenzyme F420 (Mo¨ller-Zinkhan et al. 1989; Gorris et al. 1991). The methyl group is first transferred to H4MPT and then stepwise oxidized to CO2 by the same reactions and enzymes found in methanoarchaea (> Fig. 18.2). The F420H2 formed in this oxidative pathway is reoxidized by a membrane-bound F420H2 dehydrogenase, which closely resembles the enzyme from Methanosarcina species (Kunow et al. 1994; Klenk et al. 1997). Archaeoglobus fulgidus contains a modified menaquinone, which probably functions as the electron acceptor of this

dehydrogenase. It is not yet clear how electrons are transferred from the menaquinone pool to the enzymes of sulfate reduction. Recently, a membrane-bound menaquinol-acceptor oxidoreductase that might mediate the electron transfer from the menaquinone pool to an as yet unidentified electron carrier in the cytoplasm has been isolated (Mander et al. 2002). The sequences of two of the subunits of this enzyme are related to those of the heterodisulfide reductase from Methanosarcina species, including the catalytic subunit of Hdr. However, Archaeoglobus lacks coenzymes M and B. Therefore, this heterodisulfide-reductase-like enzyme has been proposed to catalyze the reduction of an unidentified disulfide substrate, which in turn could function as an electron donor of the enzymes of sulfate reduction, such as APS reductase and sulfite reductase.

Tetrahydromethanopterin-Dependent Formaldehyde Oxidation in Methylotrophic Bacteria In the metabolism of aerobic methylotrophic bacteria, formaldehyde is formed as a central intermediate from various C1 substrates. Different pathways of formaldehyde oxidation to CO2 are known, one being tetrahydromethanopterin dependent. The H4MPT-dependent pathway was first discovered in Methylobacterium extorquens. This organism, in addition to the tetrahydrofolate-dependent pathway, has an H4MPT-dependent route for formaldehyde oxidation, which is now believed to be the main catabolic route in this organism (Chistoserdova et al. 1998). The pathway involves three H4MPT-dependent steps, which are catalyzed by an NADH-dependent methyleneH4MPT dehydrogenase, a methenyl-H4MPT cyclohydrolase, and a formyltransferase/hydrolase complex. H4MPT-dependent enzymes have also been detected in many other methylotrophic proteobacteria. For a more detailed review, see > Chap. 7, ‘‘Aerobic Methylotrophic Prokaryotes’’ in this volume.

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Physiology and Biochemistry of the Methane-Producing Archaea

F420 in Nonmethanogenic Organisms Coenzyme F420 was first discovered in methanogenic archaea. Later, coenzyme F420 was also identified in Archaeoglobus, Mycobacterium, Nocardia, Streptomyces, cyanobacteria, and some eukaryotes (Choi et al. [2001] and literature cited therein). The role of F420 in Archaeoglobus is similar to that in methanogens. Coenzyme F420 is used by Streptomyces species for tetracycline and lincomycin biosynthesis and may be used in mitomycin C biosynthesis. In Mycobacterium and Nocardia species, coenzyme F420 is used by a coenzyme F420-dependent glucose-6-phosphate dehydrogenase. Enzymes belonging to the deazaflavin class of photolyases, which are found in the green alga Scenedesmus and the cyanobacterium Synechocystis, contain 8-hydroxyazoriboflavin (also called ‘‘coenzyme F0’’). Coenzyme F420 is a derivative of coenzyme F0.

CoM-SH in Bacterial Aliphatic Epoxide Carboxylation Until 1999, methanoarchaea were the only organisms known to possess coenzyme M, which is the smallest organic cofactor found in nature. It was then discovered that coenzyme M also plays an essential role in the bacterial metabolism of short-chain epoxyalkanes, as revealed by initial studies with Xanthobacter autotrophicus and Rhodococcus rhodochrous (Allen et al. 1999). These organisms use coenzyme M as the nucleophile for the epoxide ring opening, which results in the formation of the thioether bond between CoM-SH and a 2-hydroxyalkyl residue. After oxidation to the corresponding 2-ketoalkyl-CoM intermediate, the thioether bond is attacked by a cysteine residue present in the active site of one of the key enzymes of the pathway. This results in the formation of a mixed disulfide between CoM-SH and the active-site cysteine and a carbanion, which becomes carboxylated. Reduction of the mixed disulfide in an NADH-dependent step regenerates coenzyme M. Coenzyme M seems to be ideally suited as a nucleophile and carrier molecule in this pathway (reviewed in Ensign and Allen 2003).

Do Anaerobic Methane Oxidizers Use the Methanogenic Pathway in Reverse? Although the elucidation of the pathway of CO2 reduction in methanogens required the discovery of a large number of novel coenzymes and enzymatic reactions, many of these catalysts were subsequently found in other organisms. For many years, the reaction catalyzed by methyl-coenzyme M reductase seemed to be the only step of the pathway that was truly unique to the methanoarchaea. However, very recently, genes encoding a methyl-coenzyme M reductase-like enzyme were identified in habitats where methane-oxidizing microbial communities are abundant (Hallam et al. 2003). From the biomass of one of these habitats, a methyl-coenzyme M reductase-like enzyme was isolated (Kra¨ger et al. 2003). This protein harbored a nickel-

containing prosthetic group that was identified as a heavier (mass of 951 Da) variant of coenzyme F430 (mass of 905 Da), the unique nickel porphinoid in Mcr. These studies led to the proposal that anaerobic methane oxidation biochemically, in principle, is a reversal of methanogenesis. For more details on anaerobic methane oxidation, see > Chap. 17, ‘‘Anaerobic Biodegradation of Hydrocarbons Including Methane’’ in this volume.

Regulation of Gene Expression Regulation of Catabolic Enzymes by Substrate Availability Many methanoarchaea use only one or two energy substrates so that one may not expect extensive metabolic regulation. Nevertheless, it was found that even organisms using H2/CO2 as the sole growth substrate regulate the formation of some key catabolic enzymes in response to the availability of H2. One example is the differential expression of two methyl-coenzyme M reductase isoenzymes in the Methanobacteriales and the Methanococcales (Thauer 1998). In Methanothermobacter species, isoenzyme I is encoded by the mcrBDCGA operon, and isoenzyme II is encoded by the mrtBDGA operon. The two isoenzymes differ in their catalytic properties. Isoenzyme I has a lower Vmax as compared to isoenzyme II but displays lower KM values for its substrates, CoB-SH and methyl-coenzyme M (Bonacker et al. 1993). Expression of the two isoenzymes is differently regulated by the availability of hydrogen. Isoenzyme I is predominantly formed when growth is limited by the H2 supply whereas isoenzyme II predominates when the H2 supply is not growth-rate limiting (Bonacker et al. 1992; Morgan et al. 1997). In the latter case, the methyl-coenzyme M reductase reaction might be a bottleneck of the pathway. Therefore, it could be of physiological relevance to synthesize an enzyme with a higher Vmax. There are conflicting results with respect to the regulation of other methanogenic enzymes in response to the H2 availability. Two groups found that the formation of Hmd in Methanothermobacter species parallels that of isoenzyme II of Mcr (encoded by the mrt operon), while the formation of Frh and Mtd parallels that of isoenzyme I of Mcr (encoded by the mcr operon; Morgan et al. 1997; Vermeij et al. 1997). Two other groups did not observe a formation of these enzymes in response to H2 availability with their systems (Afting et al. 2000; Luo et al. 2002). But all groups observed the same pattern of formation of McrI and McrII. The formation of flagella in Methanocaldococcus jannaschii is another example of regulation in response to H2 availability. Although flagella are not directly involved in catabolic processes, they are essential for finding optimal substrate conditions. Under H2-excess conditions, M. jannaschii cells are devoid of flagella and have almost undetectable levels of four flagellarelated proteins. Flagella synthesis occurs when H2 becomes limiting (Mukhopadhyay et al. 2000).

Physiology and Biochemistry of the Methane-Producing Archaea

Many species of hydrogenotrophic methanogens use formate in place of H2 as the electron donor for CO2 reduction. The ability to use formate is attributed to formate dehydrogenase (Fdh), which in methanoarchaea catalyzes the formatedependent reduction of coenzyme F420. The Methanococcus maripaludis genome contains two formate dehydrogenase gene clusters. The transcription of both gene clusters was found to be controlled by the availability of H2. Only in the absence of H2 was maximal expression of both fdh gene clusters observed. In contrast, formate had no marked effect on the expression (Wood et al. 2003). In contrast, expression of formate dehydrogenase in Methanobacterium formicicum seems not to be regulated (Schauer and Ferry 1980). Methanogenium thermophilum can use 2-propanol as sole electron donor for CO2 reduction. The secondary alcohol dehydrogenase responsible for 2-propanol oxidation was only formed when H2 became limiting, irrespective of the presence of the alcohol. In other methanoarchaea able to grow with secondary alcohols, formation of alcohol dehydrogenase was dependent on the availability of an alcohol, irrespective of the presence of H2 (Widdel and Wolfe 1989). Another response to H2 limitation is the synthesis of an autolytic enzyme by Methanobacterium wolfei (Kiener et al. 1987). The physiological role of this suicidal process is not known. It may be related to the induction of a defective bacteriophage (Stettler et al. 1995). The regulation of the genes encoding methanogenesis from acetate in Methanosarcina species is also well studied. Early work had already shown that acetate is only used as energy substrate when none of the higher energy-yielding substrates methanol, methylamines, or H2/CO2 are available, indicating that acetate catabolism is repressed by these other substrates (reviewed in Zinder 1993). This is consistent with the observation that the key enzymes of acetate metabolism (i.e., acetate kinase, phosphotransacetylase, acetyl-CoA synthase/carbon monoxide dehydrogenase complex, and carbonic anhydrase) are formed at a lower level in cells grown on methanol as compared to acetategrown cells (Jablonski et al. 1990). Regulation was shown to be at the mRNA level (Sowers et al. 1993; Singh-Wissmann and Ferry 1995). On the other hand, most of the enzymes necessary for the reversible reduction of CO2 to the level of methyltetrahydromethanopterin are present at a much lower level in acetate-grown cells (Jablonski et al. 1990; Mukhopadhyay et al. 1993). When Methanosarcina spp. are cultivated on methanol in the presence of H2/CO2, the oxidative branch of the methylotrophic pathway is repressed. This result is consistent with the observation that several enzymes of this pathway are formed at a lower level under these conditions (Mukhopadhyay et al. 1993). In conclusion, catabolic gene expression in Methanosarcina appears similar to systems in bacteria, which are regulated for preferential utilization of the most energetically favorable substrate. For none of the regulatory systems described above has the primary sensor and the signal transduction cascade been elucidated. However, in Methanothermobacter thermautotrophicus, studies have been performed which led to the proposal

18

that coenzyme F390 could function as a reporter compound for H2 limitation. Coenzyme F390 is formed from coenzyme F420 by adenylation or guanylation at its 8-hydroxy-group. This reaction is catalyzed by coenzyme F390 synthetase (Vermeij et al. 1994). This enzyme specifically uses oxidized coenzyme F420 as substrate, while reduced coenzyme F420 (F420H2) acts as a competitive inhibitor. Coenzyme F390 can be hydrolyzed to coenzyme F420 and AMP or GMP in a reaction catalyzed by coenzyme F390 hydrolase (Vermeij et al. 1995). This latter enzyme is redox sensitive and is inactivated by O2. Furthermore, this enzyme is activated by CoM-SH but inactivated by CoM-S-S-CoB. On the basis of the biochemical properties of these two enzymes, it has been predicted that the level of coenzyme F390 should be low when cells receive sufficient H2 (which leads to a high coenzyme F420H2 to coenzyme F420 ratio and high CoM-SH to CoM-S-S-CoB ratio). Conversely, the coenzyme F390 concentration in the cell should increase when H2 becomes limiting. This prediction was confirmed experimentally (Vermeij et al. 1997). In further studies, a Methanothermobacter thermautotrophicus mutant was isolated that was unable to grow under H2-deprived conditions. This mutant was also unable to form coenzyme F390. It also lacked the ability to synthesize isoenzyme I of Mcr, which is the enzyme preferentially synthesized under H2-limiting conditions (Pennings et al. 1998). This gives further evidence for an important role of coenzyme F390 in the response of the cell to varying H2 concentrations.

Regulation of Catabolic Enzymes by Trace Element Availability In the methanogenic pathways, enzymes containing transition metals in their active site play an essential role. Therefore, not surprisingly, these organisms have developed strategies to cope with limitations on the availability of these metal ions. One example is the synthesis of different isoenzymes of formylmethanofuran dehydrogenase (Fmd; reviewed in Vorholt and Thauer 2002). Methanothermobacter marburgensis and Methanothermobacter wolfei form two different isoenzymes, one containing tungsten bound to the molybdopterin cofactor (Fmd-W) and a second containing molybdenum bound to the molybdopterin cofactor (Fmd-M). Whereas Fmd-W is formed constitutively, Fmd-M is only formed when molybdenum is available (Hochheimer et al. 1996). A DNA-binding protein, called ‘‘Tfx,’’ was found to specifically bind to a DNA region downstream of the promoter of the fmdECB operon, which encodes Fmd-M. Therefore, Tfx may be a transcriptional regulator of the fmdECB operon (Hochheimer et al. 1999). A different set of Fmd enzymes is found in Methanopyrus kandleri. This organism forms two tungsten-containing Fmd isoenzymes (Vorholt et al. 1997). One isoenzyme (called ‘‘Fwu’’) contains selenium, whereas the second (called ‘‘Fwc’’) does not. In general, Fmd contains a conserved cysteine residue, which is also conserved in other molybdopterin-containing

655

656

18

Physiology and Biochemistry of the Methane-Producing Archaea

enzymes. From the crystal structure of other molybdopterincontaining enzymes, for example, dimethylsulfoxide reductase, this residue is known to provide a ligand to the molybdenum center. In Fwu, this cysteine residue is replaced by selenocysteine. The gene encoding the catalytic subunit FwuB is in the polycistronic operon fwuGDB. The gene encoding FwcB, the catalytic subunit of Fwc, is transcribed monocistronically. During growth of the organism on medium supplemented with selenium, only the fwuGDB operon is transcribed. During growth under selenium limitation, both fwuGDB and fwcB are transcribed. Selenium-dependent gene expression has also been observed in Methanococcus voltae. In this organism, two isoenzymes of the coenzyme F420-reducing hydrogenase (called ‘‘Fru’’ and ‘‘Frc’’) and two isoenzymes of the coenzyme F420-nonreducing hydrogenase (called ‘‘Vhu’’ and ‘‘Vhc’’) are encoded in the genome (Sorgenfrei et al. 1997). One enzyme of each type, Fru and Vhu, contains selenocysteine in the hydrogen-activating reactive site. The corresponding isoenzymes, Frc and Vhc, have a cysteinyl residue in the homologous positions. The two selenium-containing hydrogenases are constitutively expressed. The operons vhc and frc encoding the selenium-free enzymes are only transcribed under selenium limitation. They are connected by a common intergenic region comprising both promoters and positive and negative regulatory sequence elements, which were defined by mutational analyses employing a reporter gene system (Noll et al. 1999). A putative activator protein has been identified but not yet further characterized (Mu¨ller and Klein 2001). A protein binding to a negative regulatory element involved in the regulation of the two operons was purified. Through the identification of the corresponding gene, the protein was found to be a LysR-type regulator. It was named ‘‘HrsM’’ (hydrogenase gene regulator, selenium dependent in M. voltae). Also, hrsM knockout mutants constitutively transcribed the vhc and frc operons in the presence of selenium (Sun and Klein 2004). Nickel is an essential trace element for methanoarchaea. Studies with Methanothermobacter marburgensis have shown that this organism has developed a strategy to spare nickel under nickel limitation. As outlined above, coenzyme F420-reducing hydrogenase (Frh), which is a [NiFe] hydrogenase, can be functionally replaced by the combined action of Hmd and Mtd. These two latter enzymes do not contain Ni. When M. marburgensis was cultivated under nickel-limited conditions, the specific activity of Hmd and Mtd was six- and fourfold higher and that of Frh up to 180-fold lower than in cells grown on nickel-sufficient medium. The frh transcripts were no longer detectable in cells grown under Ni limitation, whereas the relative abundance of the hmd and mtd transcripts increased (Afting et al. 1998, 2000).

Regulation of Nitrogen Assimilation Nitrogen assimilation by Methanococcus maripaludis is highly regulated. This organism fixes N2 but can also use ammonia or

alanine as sole nitrogen sources. In the presence of ammonia or alanine, N2 fixation is highly repressed (Cohen-Kupiec et al. 1997; Lie and Leigh 2002). The repressor has been isolated and is very unusual for this class of proteins. Called ‘‘NrpR,’’ it possesses very low sequence similarity to previously described DNA-binding proteins in the prokaryotes (Lie and Leigh 2003). NrpR also regulates the expression of glnA in M. maripaludis. In addition to transcriptional regulation, N2 fixation is also regulated by a switch-off mechanism. Upon the addition of ammonia or alanine, nitrogen fixation ceases immediately (Kessler et al. 2001; Lie and Leigh 2002). This regulation requires the participation of two GlnB homologues encoded by nifI1 and nifI2. Although this system acts very similarly to the bacterial system for the posttranslational ADP-ribosylation of the nitrogenase reductase, its mechanism of action is not currently known.

Bioenergetics of Growth Coupling Sites in Methanogenesis Energy conservation by methanoarchaea is via electron transport phosphorylation as outlined above. The H2/CO2 pathway contains two energy-coupling sites: the H4MPT:coenzyme M methyltransferase reaction and the reduction of the heterodisulfide. While the methyltransferase reaction is coupled to the primary extrusion of Na+, the heterodisulfide reductase reaction is coupled to the extrusion of H+. Experimental proof that the latter reaction is coupled to energy conservation is, however, only available for Methanosarcina species. Via a Na+/H+ antiporter, D:mNa+ and D:mH+ are interconvertible (Kaesler and Scho¨nheit 1989). Part of the energy conserved in these ion gradients is used to drive the reduction of CO2 to formylmethanofuran by reversed electron transport, while the remaining part of the energy is used for the synthesis of ATP via ATP synthase. Methanoarchaea contain A1A0 ATP synthases characteristic for archaea (Mo¨ller 2004). In M. thermautotrophicus and Methanosarcina mazei, this is the only ATP synthase encoded in the genome sequences. In contrast, the genomes of M. barkeri and M. acetivorans encode both (an A1A0 ATP synthase and a F1F0 ATP synthase). Expression of the latter enzyme in M. barkeri could, however, not be demonstrated (Mo¨ller 2004). The ion specificity of A1A0 ATP synthases is not yet established. In silico analysis of the proteolipid of some A1A0 ATP synthases reveal the presence of a Na+ binding motif and suggest that these enzymes use Na+ as coupling ion (Mo¨ller 2004). In aceticlastic methanogenesis, the methyltransferase and the heterodisulfide reductase reactions are also sites of energy conservation. Formation of H2 from reduced ferredoxin, catalyzed by Ech hydrogenase, might represent an additional energy-coupling site (> Fig. 18.14). On the other hand, activation of acetate to acetyl-CoA requires at least one ATP in Methanosarcina spp. and two ATP in Methanosaeta spp. Thus, cells must recover the high cost of acetate activation.

Physiology and Biochemistry of the Methane-Producing Archaea

When methanol or methylamines are used as energy substrates, the heterodisulfide reductase reaction is also a site of energy conservation. However, the H4MPT:coenzyme M methyltransferase and the formylmethanofuran dehydrogenase reactions now operate in reverse. Thus, the methyltransferase reaction becomes energy consuming while the oxidation of formylmethanofuran to CO2 and methanofuran is coupled to energy conservation.

Growth Yields Methanoarchaea possess specialized systems to generate the energy needed for growth from the process of methanogenesis, and they have only a limited capacity to metabolize complex carbon compounds. Even the secondary alcohols, which can serve as electron donors for CO2 reduction in some species, are only partially oxidized to ketones. About half of the described species of methanogens are capable of autotrophic growth and obtain all of their cellular carbon from CO2. While the remainder may require organic compounds for growth, these compounds are assimilated into cellular carbon and not extensively metabolized. Compounds typically assimilated include acetate and the volatile fatty acids like isovalerate, 2-methylbutyrate, isobutyrate, and propionate, which are common in anaerobic environments, as well as amino acids. The inability to assimilate complex organic compounds has profound effects on the energy requirements for growth. On the basis of biosynthetic pathways known and inferred from the genomic sequence, Methanococcus maripaludis, a typical hydrogenotrophic methanogen, must expend 89 mmol of ATP equivalents and 97 mmol of [2H] for the biosynthesis of a gram of cells from CO2 (> Table 18.4). Given that 50 % of the cell is carbon, the amount of reductant required is close to 84 mmol of

. Table 18.4 Bioenergetic requirements for monomer biosynthesis during growth of methanogens in mineral and rich media Requirement (mmol/g of cell dry wt.)

a

Growth conditions

Pa

[2H]b

Total [2H]c

Autotrophic growth in mineral medium + acetate

89

97

451

89

34

388

Rich mediumd

63

25

276

E. coli minimal medium

21

18

–

ATP equivalents required Reductant as NADH or H2 equivalents required for anabolism c Includes the H2 necessary for methanogenesis to make ATP with a stoichiometry of 1 ATP/CH4 d Includes acetate + the volatile fatty acids for branched-chain amino acid biosynthesis + aryl acids for aromatic amino acid biosynthesis + the nucleobases (guanine, adenine, and uracil) commonly taken up by the salvage pathway b

18

[2H], or the theoretical amount necessary to reduce 42 mmol of CO2 to the oxidation state of carbon in the cell. Presumably, the difference is due to oxidations that occur during biosynthesis and the approximation of the cell composition. The ATP requirement greatly exceeds that of a typical heterotroph such as E. coli growing in a minimal medium. It is also much larger than the approximately 36 mmol ATP (gram of cells) 1 required for polymerization reactions, which includes protein, DNA and RNA biosynthesis (Forrest and Walker 1971; Ingraham et al. 1983). Thus, monomer biosynthesis is the major energy demand for growth of a hydrogenotrophic methanogen, and the assimilation of organic carbon sources may have large effects on their growth. Many methanogens assimilate exogenous acetate, which is frequently abundant in anaerobic habitats. From the biosynthetic pathways, about 16 mmol of acetyl-CoA are utilized in the biosynthesis of one gram of cells; hence, acetate has the potential of providing about 75 % of the cellular carbon. Assuming that two ATPs are consumed to active acetate via the high affinity acetyl-CoA synthetase reaction, there is no savings in the ATP requirement for growth when compared with CO2 fixation (> Table 18.4). If only one ATP is utilized to activate acetate via the low affinity acetate kinase reaction, about 16 mmol of ATP is spared, which is about 18 % of the total ATP requirement for monomer biosynthesis. Similarly, methanogens frequently assimilate the branched-chain volatile fatty acids as sources of branched-chain amino acids and aryl acids as a source of aromatic amino acids. Together, these amino acids account for about 25 % of the cellular carbon. Assuming that the carboxylic acids are assimilated by an acyl-CoA synthetase reaction requiring two ATP equivalents, followed by ferredoxindependent oxidoreductase requiring one ATP equivalent to activate the reductant and one [2H], and an aminotransferase (which requires one ATP and one [2H] to make glutamate), four ATP equivalents and two [2H] are required for each amino acid biosynthesized. Even then, this pathway results in a large reduction in the energy requirements for growth (> Table 18.4). The maximum cell yields can be estimated. For a hydrogenotrophic methanogen fixing CO2 as its major carbon source, about 89 and 36 mmol of ATP per gram of cells are required for monomer biosynthesis and polymerization reactions, respectively. Thus, the maximal cell yield is expected to be about 8.0 g of cell dry weight per mol of ATP. For a hydrogenotrophic methanogen obtaining carbon from acetate, the volatile fatty acids and aryl acids, the yield is about 10 g of cell dry weight per mol of ATP. In contrast, for a heterotroph, the maximal cell yield is 28 g of cell dry weight per mol of ATP. For an autotroph using the Calvin cycle of CO2 fixation, the maximal cell yield is 4.75 g of cell dry weight per mol of ATP (Forrest and Walker 1971). Thus, while the cell yield of an autotrophic methanogen is considerably less than that of a heterotroph, it theoretically could be nearly twice that of a chemolithotroph using the Calvin cycle. For comparison, the observed cell yield for methanogens are usually in the range of 1–6 g of cells per mol of methane, and the measured maximal cell yields are 3–6 g of cells per mol of

657

658

18

Physiology and Biochemistry of the Methane-Producing Archaea

. Table 18.5 Genomic sequences of methanoarchaea Genome size (kbp)

Number of ORFs

Comments

Reference(s)

Methanothermobacter thermautotrophicus )H

1,751

1,855

Thermophilic hydrogenotroph

Smith et al. (1997)

Methanocaldococcus jannaschii JAL-1

1,723

1,726

Hyperthermophilic hydrogenotroph

Bult et al. (1996)

Methanococcoides burtonii

2,668

2,676

Partial sequence, psychrotolerant methylotroph

Saunders et al. (2003)

Organism

Methanococcus maripaludis S2

1,661

1,722

Mesophilic hydrogenotroph

Hendrickson et al. (2004)

Methanogenium frigidum

1,598

1,815

Partial sequence, psychrophilic hydrogenotroph

Saunders et al. (2003)

Methanopyrus kandleri AV19

1,695

1,692

Hyperthermophilic hydrogenotroph

Slesarev et al. (2002)

Methanosarcina acetivorans C2A

5,751

4,524

Mesophilic acetotroph and methylotroph

Galagan et al. (2002)

Methanosarcina barkeri Fusaro

4,830

5,066

Partial, mesophilic acetotroph and methylotroph

Joint Genome Institute, unpublished

Methanosarcina mazei Go¨1

4,096

3,371

Mesophilic acetotroph and methylotroph

Deppenmeier et al. (2002)

methane (Vogels et al. 1988; Tsao et al. 1994; Schill et al. 1996). In the literature, the Methanosarcina species appear to have higher growth yields on H2/CO2 than Methanothermobacter species and others, but these results are from different laboratories and observed under different growth conditions. Following cultivation of two mesophiles, Methanosarcina barkeri and Methanobrevibacter aboriphilus, under the similar conditions on H2/CO2, the cell yields were 4.2 g and 1.4 g of dry cells per mol of CH4, respectively (R. Hedderich, unpublished data). These results confirmed the lower cell yield among the Methanobacteriales. Possibly, the lower cell yield might result from a different mechanism of coupling methanogenesis to the proton motive force or from higher maintenance energy during growth.

Genomes of the Methanoarchaea The complete genomes have been sequenced in a representative of every order of the methanoarchaea except the Methanomicrobiales, where only a partial sequence is available (> Table 18.5). The sizes of the genomes vary from 1.6 to 5.8 Mbp, reflecting the great diversity in this group of organisms (> Table 18.5). In general, the genomes of the hydrogenotrophic methanogens are smaller, in the range of 1.6–1.8 Mbp. Even among these small genomes, the gene content is not highly conserved, and only about two-thirds of the genes in any one organism are likely to be conserved within the methanoarchaea (W. B. Whitman, unpublished observation). The genomes of the methylotrophic methanogens are much larger, in the range of 2.7–5.8 Mbp. In the Methanosarcina spp., the large genome

seems to have followed the acquisition of a large number of genes from the anaerobic Firmicutes (Deppenmeier et al. 2002) and may be responsible in part for the wide substrate specificity of these organisms (Galagan et al. 2002). The growth temperature optima of those methanoarchaea whose genomic sequences have been determined are 15–98  C. Thus, it has been possible to make detailed correlations of certain structural features in proteins and nucleic acids with growth temperature (Saunders et al. 2003). The amino acid leucine was highly enriched in the proteins from organisms with high growth temperatures, while the amino acids glutamine and threonine were highly enriched at low growth temperatures. In addition, the proteins of organisms with high growth temperatures were enriched in the mean fraction of charged residues in the solvent accessible as well as solvent inaccessible areas. Likewise, the contribution of hydrophobic residues to the solvent accessible area decreased with growth temperature. The tRNAs of organisms with high growth temperatures also possessed higher mol% G + C contents, especially in the stem regions (Saunders et al. 2003).

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