Handbook of Hydrocarbon and Lipid Microbiology [1 ed.] 9783540775843, 9783540775874

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Handbook of Hydrocarbon and Lipid Microbiology

Kenneth N. Timmis

Handbook of Hydrocarbon and Lipid Microbiology

With 712 Figures* and 277 Tables

*For color figures please see our Electronic Reference on www.springerlink.com

Professor Kenneth N. Timmis Environmental Microbiology Laboratory Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany

Library of Congress Control Number: 2009930947 ISBN: 978–3–540–77584–3 This publication is available also as: Electronic publication under ISBN 978–3–540–77587–4 and Print and electronic bundle under ISBN 978–3–540–77588–1 This work is subject to copyright. All rights are reserved, 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 other ways, and storage in data banks. Duplication of this publication or parts thereof is only permitted under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer‐Verlag. Violations are liable for prosecution under the German Copyright Law. ß Springer‐Verlag Berlin Heidelberg 2010 The use of registered names, trademarks, 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. Springer is part of Springer Science+Business Media springer.com Editor: Christina Eckey/Sandra Fabiani, Heidelberg, Germany Development Editor: Sylvia Blago, Heidelberg, Germany Production: SPI‐Publishing, Pondicherry, India Cover Design: Frido Steinen‐Broo, Girona, Spain Printed on acid‐free paper

SPIN: 12212507

2109 — 5 4 3 2 1 0

This Handbook is dedicated to the three waves of hydrocarbon and lipid microbiology and biochemistry researchers. Firstly, the early pioneers, who were the first to be fascinated by the unique biology and discovery potential of the topic, solved basic technical challenges, made seminal discoveries, mapped out the field, laid a solid foundation for the current explosion of exciting research, and acted as role models for successive generations of scholars. Secondly, the future pioneers, the brightest students in the labs of the authors of this Handbook and of others in the field who represent the new generation of ‘‘greasy’’ microbiologists: may this Handbook inspire them to tenaciously explore thus far undiscovered facets of biological activities at hydrophobic:hydrophilic substance interfaces, to seek the causes and consequences of molecular domains/molecules/multimolecular structures skating on and slipping in and out of hydrophobic surfaces, to find answers to the most challenging questions, to surmount the most difficult technical hurdles, and to illuminate the ecophysiological controls that interlink and intermodulate apparently unrelated diverse activities at all scales and that integrate such activities with environmental parameters. Most importantly may they have much intellectual excitement and fun while advancing the knowledge frontiers of this critically important and fascinating subject. And thirdly, the current generation of greasy microbiologists, especially those who sacrificed their valuable research and family time to write chapters for this Handbook, and particularly those authors who accepted our invitations in the knowledge that they were pregnant. To research, formulate and edit chapters during a period of gestation, delivery and nurturing of a baby, and dealing with the incompatibility of creativity and lack of sleep (holding the baby with one hand and clicking, scrolling, typing with the other to some extent compromises both activities; hopefully the night feeds were without computer and thus allowed bonding catch-up, and that the imprinting process will not result in a new generation of future mothers who can only feed while typing manuscripts and muttering about greasy microbes, demands a very special quality, not to mention quantity, of motivation: thank you! This Handbook is gratefully dedicated to you all.

Preface

‘‘Water is life!’’ All active cellular systems require water as the principal medium and solvent of their metabolic and ecophysiological activities. Hydrophobic compounds and structures, which tend to exclude water, though providing inter alia excellent sources of energy and a means of biological compartmentalization, present problems of cellular handling, poor bioavailability and, in some cases, toxicity. Microbes both synthesize and exploit a vast range of hydrophobic organics, especially petroleum oil hydrocarbons and industrial pollutants, and the underlying interactions not only have major consequences for the lifestyles of the microbes involved, but also for biogeochemistry, climate change, environmental pollution, human health and a range of biotechnological applications. In this Handbook we attempt to cover the exceptional range of cellular, population and community activities of microbes interacting with the major hydrophobic organics found in the biosphere and to encapsulate the consequences of such interactions for the health and disease of microbes, higher organisms and the environment, as well as the beneficial applications that have been or can be developed therefrom. In order to achieve this goal with a text of reasonable length, it was decided to conceive a Handbook consisting of many short chapters dealing with their topics in a relatively superficial manner, while citing original references that document the principal advances described, rather than of long chapters providing detailed, comprehensive coverage of individual topics, some of which already exist in other excellent texts. Although this Handbook started with a game plan, a framework of topics to be covered, it evolved continuously throughout the chapter recruitment period, as more and more new facets of microbe:hydrocarbon/lipid interactions became apparent, many of which were entirely new at the time of recruitment, which underscores both the explosion of highly original research in the area, and the continuous discovery of new activities and consequences of microbial activities at hydrophobic:hydrophilic substance interfaces. Eventually, in order to move the Handbook into production (and to avoid a revolution among authors who had provided chapters on time), we reluctantly had to stop adding new topics. During the process of literature review, chapter conception, text formulation, and chapter reviewing and editing, authors and editors alike became exposed to exciting new information that, in some cases, sharply increased creativity. This is of course a major motivation for participation in such an effort. As far as we are aware, this Handbook breaks new ground in attempting to deal generically with microbial interactions with major hydrophobic organics in the biosphere, that is, with the fundamental problem of hydrophobicity for water-based cellular systems, and in exploring generically the wide range of consequences for (wo)mankind of such interactions. Volume 1 sets the scene for hydrophobic microbiology with introductory chapters that describe the main hydrophobic organic compounds in the biosphere, their diversity and physico-chemical characteristics, how they are formed, how they enter the biosphere (including via anthropogenic activities like mining, accidents and illegal dumping), and where in the biosphere

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Preface

they are to be found. Subsequent chapters deal with the biogenesis of hydrophobic compounds and organelles - methane, other hydrocarbons, fatty acids, lipids, oils and wax esters - their biochemistry, genetics, genomics, microbiology and ecophysiology. The second Volume constitutes a sequel to the production of hydrophobic compounds, and deals with their consumption. It not only outlines the biochemistry, genetics and ecology of utilization, but also describes the physiological problems of interactions with such compounds, like poor bioavailability, uptake, toxicity and problems of an acetyl-CoA-centric metabolism. Following this framework of the generic biology underlying microbe:oil interactions, Volume 3 provides upto-date descriptions of the biological players in these interactions: the hydrocarbon-utilizing microbes themselves and the microbial communities in which they function. With the fundamentals of hydrocarbon and lipid production and consumption by microbes documented, Volume 4 is dedicated to the wide range of consequences of these interactions. These include hydrocarbon biodegradation and bioremediation, biomonitoring, fuel production, chemicals production, global consequences, and consequences for the health and disease of higher organisms. Volume 4 ends with a brief selection of author predictions on the future of ‘‘greasy microbiology’’. To complete the Handbook, Volume 5 provides a comprehensive set of methods and protocols for research in greasy microbiology. This is to our knowledge the first and only, much needed collection of state-of-the-art protocols for research in this topic and will, we anticipate, become a major experimental resource for the community that facilitates and accelerates important advances. Volume 5 ends with an Annex of literature and Web resources pertinent to the topic of hydrocarbon and lipid microbiology. In addition to dealing with the generic biology of microbial interactions with hydrophobic substances, the Handbook comprises a unique combination of topic overviews that includes:

– most up-to-date listing and description of hydrocarbon-producing and -consuming microbes

– ecophysiology of hydrocarbon-producing and -consuming microbial communities in the diverse hydrocarbon-containing habitats of the biosphere

– global overview of key metabolic routes to and from central metabolism from and to hydrocarbons, lipids and hydrophobic storage products.

– enzymology/catalytic mechanisms underlying HC-processing in aerobic and anaerobic – – – – – – – –

environments genetics of relevant pathways and genomics of key model strains stresses imposed by hydrocarbons, physiological responses and tolerance mechanisms hydrocarbon microbiology and global warming the range of applications of hydrocarbon-metabolizing microbes alternative energy sources oleochemical biotechnology enzyme and metabolic engineering; systems biology of industrial processes hydrocarbons, lipids and microbial infections of humans, animals and plants

In selecting topics to be covered, and the chapters and authors needed to do these topics justice, we were supported by a blue-blooded Scientific Advisory Board of leading researchers in the various sectors of ‘‘greasy’’ microbiology. These experts not only helped frame the contents of the Handbook and fingered potential authors of chapters, but also in most cases contributed one or more chapters and carried a considerable burden of the reviewing of chapters by others. Following a gentle but firm suggestion from my very good friend, Victor de Lorenzo, that I delegate part of the editorial load, a subgroup of the SAB subsequently accepted

Preface

to take on editorial responsibilities, resulting in the Handbook acquiring three Section Editors: Terry McGenity, for taxonomic sections, Jan Roelof van der Meer, for the methods sections, and Victor de Lorenzo, for genetic/genomic sections. I am most grateful to Victor for his advice, and to him, Terry and Jan Roelof for the super efforts they invested in the Handbook. They were fun people with which to work on this project. In addition to the SAB, a number of contributors to the Handbook provided enthusiastic support over and above the call of duty, in terms of advice, author and topic suggestions, and offers to cover topics lacking authors. These include, in no particular order, Heinz Wilkes, Otto Geiger, Karin Athenstaedt, Dietmar Pieper, Hauke Harms, Lukas Wick, Ibrahim Banat, Becky Parales, Herman Heipieper, Julia Foght, Sylvie Le Borne, Antje Boetius, Charles Greer, Eugene Rosenberg, Manolo Ferrer, Karl-Erich Ja¨ger, Monica Bassas, Sagrario Arias, Ann Wood, and the FaceIt consortium. Finally, I wish to express my undying gratitude to the people who not only made this major work possible, but also made it such a satisfying project:

– the Section Editors, Victor, Terry and Jan Roelof, who enthusiastically dedicated much

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time, effort and creative energy to the project, including the writing of (additional) chapters they thought were important that failed to be delivered by authors who had initially accepted, the SAB, who supported and advised during the conception and realization of the Handbook, and reviewed many of the manuscripts, some special people - Don Kelly and Ann Wood, Willy Verstraete, Fritz Widdel, Sylvie Le Borne, Roger Prince, Dietmar Pieper, Misha Yakimov and Colin Murrell - who became truly infected with the excitement of the project and committed themselves to so many chapters, some proposed to them, some they proposed themselves, and whose enthusiasm raised the fun level of the project to unanticipated heights, the folks at Springer, especially Christina Eckey, who initiated the whole thing, Lydia Mu¨ller, Simone Giesler and Sylvia Blago, who were the production interface with authors and editors and did all the manuscript processing, and Dieter Czeschlik, the senior figure in the publication of this tome. I have had a number of editorial experiences during a long career, but this was certainly the most enjoyable, as a result of the enthusiasm, responsiveness, friendliness and sense of humour of these professionals. and, finally, my long term wife and partner, Joan, and our son, James, who also sacrificed family quality time in the cause of the Handbook and, in the case of Joan, spent long hours on the internet seeking and re-seeking authors’ affiliations and e-mail addresses, and learning that the web is as chaotic as the Editor of this Handbook.

Thank you all for your exceptional efforts, enthusiasm and support in creating a work summarizing exciting discoveries and impressive research achievements in hydrocarbon microbiology that we all hope will focus the spotlight on an exciting group of interconnected biological research topics that have major implications for society, its energy supplies, chemicals and pharma, nutrition, health and disease, the environment, global warming, and their future development, and that will stimulate and inspire new research efforts and the exploration of new directions. Kenneth Timmis HZI, Braunschweig, April 2009

ix

About the Editor Kenneth N. Timmis Environmental Microbiology Laboratory Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany [email protected]

Kenneth Timmis studied microbiology and obtained his Ph.D. at Bristol University. He undertook post-doctoral training at the Ruhr-University Bochum, Yale and Stanford, at the latter two as a Fellow of the Helen Hay Whitney Foundation. He was then appointed Head of an Independent Research Group at the Max Planck Institute for Molecular Genetics in Berlin, then Professor of Biochemistry in the University of Geneva, Faculty of Medicine. Thereafter, for almost 20 years, he was Director of the Division of Microbiology at the National Research Centre for Biotechnology (GBF)/now the Helmholtz Centre for Infection Research (HZI). He is currently Head of the Environmental Microbiology Laboratory at the HZI and Professor of Microbiology at the Technical University, Braunschweig. Kenneth Timmis has worked for more than 30 years in the area of environmental microbiology and biotechnology, has published over 400 peer-reviewed original research papers in international journals, and is an ISI Highly Cited Microbiology-100 researcher. His group has worked for many years, inter alia, on the biodegradation of oil hydrocarbons, especially the genetics and regulation of toluene degradation, pioneered the topic of experimental evolution of novel catabolic activities, discovered the new group of marine hydrocarbonoclastic bacteria, and initiated genome sequencing projects on bacteria that are paradigms of microbes that degrade organic compounds (Pseudomonas putida and Alcanivorax borkumensis). He is Fellow of the Royal Society, Member of the EMBO, Fellow of the American Academy of Microbiology, Member of the European Academy of Microbiology, Recipient of the Erwin Schro¨dinger Prize, and Scientific Advisory Board Member of leading research institutes. He founded the journals Environmental Microbiology and Microbial Biotechnology.

Section Editors Terry McGenity Department of Biological Sciences University of Essex Wivenhoe Park CO4 3SQ Colchester United Kingdom [email protected]

Terry McGenity is a senior lecturer at the University of Essex, UK. His PhD, investigating the microbial ecology of ancient salt deposits (University of Leicester), was followed by postdoctoral positions at the Japan Marine Science and Technology Centre (JAMSTEC, Yokosuka) and the Postgraduate Research Institute for Sedimentology (University of Reading). He worked as a postdoc with Ken Timmis at the University of Essex, where he was inspired to investigate microbial interactions with oil at multiple scales, from communities to cells, and as both a source of food and stress. He has broad interests in microbial ecology and diversity, particularly with respect to carbon cycling, and is driven to better understand how microbes cope with, or flourish in, hypersaline, desiccated and poly-extreme environments.

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Section Editors

Jan Roelof van der Meer Department of Fundamental Microbiology University of Lausanne Baˆtiment Biophore, Quartier UNIL-Sorge 1015 Lausanne Switzerland [email protected]

Jan Roelof van der Meer is Professor of Environmental Microbiology at the University of Lausanne, Switzerland. The main focus of his current research is the evolutionary adaptation of bacteria to hydrocarbon pollution, and the use of bacteria as microscale sensors for hydrocarbon pollution and bioavailability. He studied Environmental Sciences at, and received his Ph.D. in 1992 from, the University of Wageningen in The Netherlands. From 1992 to 2003 he worked as Group Leader at the Swiss Federal Institute of Aquatic Sciences (EAWAG) near Zu¨rich. He is currently coordinating the sixth Framework STREP Project FaceIt, and the seventh Framework Project BACSIN, which assembled key groups in Europe to work on pollution effects and biodegradation. One of the goals of the FaceIt project, in which Victor de Lorenzo, Terry McGenity and Ken Timmis were also involved, was to assemble and publish a collection of methods and strategies for laboratory and field work in hydrocarbon microbiology. This aim ultimately merged with the concept of the Handbook to produce Volume 5. We are very grateful for the invaluable help, time, energy, and expertise that the FaceIt project partners, along with the many other contributors, have invested in this volume.

Section Editors

Victor de Lorenzo Systems Biology Program Centro Nacional de Biotecnologı´a-CSIC Campus de Cantoblanco 28049 Madrid Spain [email protected]

Victor de Lorenzo (Madrid, 1957) is currently Professor of Research at the Spanish National Centre for Biotechnology. Although he is a chemist by training, he developed an interest for Microbiology and Environmental Biotechnology. After his PhD in Madrid (1993, Autonomous University) and various postdoctoral positions in Paris (1984, Institut Pasteur), Berkeley (1985–1987, University of California), Geneva (1988, Centre Medical Universitaire), and Braunschweig (1989–1990, Gesellschaft fur Biotechnologische Forschung), he decided to pursue the problem of environmental pollution, not only as a biotechnological challenge, but also as the source of fundamental questions in biology. These include the evolutionary emergence of catalytic functions and their regulatory proteins and networks, as well as the mechanisms behind the metabolic economy of cells under natural conditions. Since his postdoc with Ken Timmis in the GBF in the late 1980s, he developed a taste for the design of genetic tools that continues to this day. Many such tools are used by hundreds of laboratories around the world, and many others are in the pipeline for facilitating the application of synthetic biology to environmental questions.

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Table of Contents

Dedication . . . . . . . Preface . . . . . . . . . . About the Editor . . . Section Editors . . . . List of Contributors

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.. v . . vii . . xi . xiii . xlix

Volume 1 Hydrocarbons, Oils and Lipids: Diversity, Properties and Formation Part 1 Diversity and Physico-Chemical Characteristics . . . . . . . . . . . . . . . . . . . . . 1 1

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 H. Wilkes . J. Schwarzbauer

2

Methods of Hydrocarbon Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 H. Wilkes

3

Natural Gas Hydrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 J. M. Schicks

4

Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes: Analytical Methods, Diversity, Characteristics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 79 W.-R. Abraham

Part 2 Formation and Location . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 5

Stable Isotopes in Understanding Origin and Degradation Processes of Petroleum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 A. Vieth . H. Wilkes

6

The Microbial Production of Methane and Other Volatile Hydrocarbons . . . 113 M. Formolo

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7

Isoprene, Isoprenoids and Sterols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 127 J. Harder

8

Hopanoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 M. Rohmer

9

Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 A. Pearson

10

Global Relations Between the Redox Cycles of Carbon, Iron, and Sulfur . . . . 157 W. E. Krumbein . A. Gorbushina

11

History of Life from the Hydrocarbon Fossil Record . . . . . . . . . . . . . . . . . . 171 C. C. Walters . K. E. Peters . J. M. Moldowan

Part 3 Transfer from the Geosphere to Biosphere . . . . . . . . . . . . . . . . . . . . . . . 185 12

Marine Cold Seeps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 E. Suess

13

Mud Volcanoes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 H. Niemann . A. Boetius

14

Abiogenic Hydrocarbon Production at the Geosphere-Biosphere Interface via Serpentinization Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . 215 G. Proskurowski

15

Coking Processes and Manufactured Gas Plants and Their Environmental Impact on Soil and Groundwater . . . . . . . . . . . . . . . . . . . . 233 N. Hu¨sers . P. Werner

16

Shipping-Related Accidental and Deliberate Release into the Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 243 C. Gertler . M. M. Yakimov . M. C. Malpass . P. N. Golyshin

17

Oil Tanker Sludges and Slops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 257 C. Gertler . M. M. Yakimov . M. C. Malpass . P. N. Golyshin

Part 4 Environmental Chemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267 18

Chemistry of Volatile Organic Compounds in the Atmosphere . . . . . . . . . . 269 R. Koppmann

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19

Hydrocarbons in the Pedosphere . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 279 L. Schwark

20

Organic Matter in the Hydrosphere . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 297 J. Schwarzbauer

Part 5 Biochemistry of Biogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 319 21

Introduction to Microbial Hydrocarbon Production: Bioenergetics . . . . . . . 321 M. J. McInerney . T. Hoehler . R. P. Gunsalus . B. Schink

22

Methanogenesis: Syntrophic Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . 337 J. R. Sieber . M. J. McInerney . C. M. Plugge . B. Schink . R. P. Gunsalus

23

Biochemistry of Acetotrophic Methanogenesis . . . . . . . . . . . . . . . . . . . . . . 357 J. G. Ferry

24

Aliphatic Hydrocarbons, Carbon–Carbon Bond Formation . . . . . . . . . . . . . 369 L. P. Wackett

25

Halogenated Organic Compounds – Carbon-Halogen Bond Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 375 C. D. Murphy

26

Formation of Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 385 I. M. Lo´pez-Lara . O. Geiger

27

Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 395 O. Geiger . C. Sohlenkamp . I. M. Lo´pez-Lara

28

Lipid A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 409 R. E. Bishop

29

Membrane Biogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 417 H. Goldfine

30

Membrane Disrupting Proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425 J. H. Lakey . G. Anderluh

31

Lipid Intermediates in Bacterial Peptidoglycan Biosynthesis . . . . . . . . . . . 435 J. van Heijenoort

32

Phenolic Lipids Synthesized by Type III Polyketide Synthases . . . . . . . . . . 445 A. Miyanaga . S. Horinouchi

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33

The Biosynthesis and Evolution of Archaeal Membranes and Ether Phospholipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 451 Y. Koga

34

Production of Wax Esters by Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 459 J.-F. Rontani

35

Neutral Lipids in Yeast: Synthesis, Storage and Degradation . . . . . . . . . . . 471 K. Athenstaedt

Part 6 Genetics of Biogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 481 36

Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 483 M. Rother

37

Functional Genomics of Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . 501 B. Lupa

38

Regulation of Membrane Lipid Homeostasis in Bacteria . . . . . . . . . . . . . . . 509 M. A. Martinez . G. E. Schujman . H. C. Gramajo . D. de Mendoza

39

Type III Polyketide Synthases Responsible for Phenolic Lipid Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 519 A. Miyanaga . S. Horinouchi

40

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria . . . . . . 527 R. Kalscheuer

41

Players in the Neutral Lipid Game – Proteins Involved in Neutral Lipid Metabolism in Yeast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 537 K. Athenstaedt

Part 7 The Microbes (Section Editor: Terry McGenity) . . . . . . . . . . . . . . . . . . . . . 547 42

Taxonomy of Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 549 Y. Liu

43

Methanobacteriales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 559 Y. Liu

44

Methanococcales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 573 Y. Liu

45

Methanomicrobiales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 583 Y. Liu

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46

Methanosarcinales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 595 Y. Liu

47

Methanopyrales . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 605 Y. Liu

48

Aliphatic Hydrocarbon Producers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 609 L. P. Wackett

Part 8 Methanogenic Communities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 615 49

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 617 O. R. Kotsyurbenko

50

Soil, Wetlands, Peat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 625 O. R. Kotsyurbenko

51

Environmental Constraints that Limit Methanogenesis . . . . . . . . . . . . . . . 635 T. Hoehler . R. P. Gunsalus . M. J. McInerney

52

Methanogenesis in Arctic Permafrost Habitats . . . . . . . . . . . . . . . . . . . . . . 655 D. Wagner . S. Liebner

53

Methanogens and Methanogenesis in Hypersaline Environments . . . . . . . 665 T. J. McGenity

54

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems . . . . . . . . . . . . . 681 B. Ollivier . J.-L. Cayol

55

Mammalian Digestive Tract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 693 G. N. Jarvis . D. Al-Halbouni

56

Methanogenesis in the Digestive Tracts of Insects . . . . . . . . . . . . . . . . . . . 707 A. Brune

Volume 2 Microbial Utilization of Hydrocarbons, Oils and Lipids Part 1 Introduction: Theoretical Considerations . . . . . . . . . . . . . . . . . . . . . . . . . 729 1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 731 F. Widdel . F. Musat

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Part 2 Biochemistry of Aerobic Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . 765 2

Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 767 T. J. Smith . Y. A. Trotsenko . J. C. Murrell

3

Enzymes for Aerobic Degradation of Alkanes . . . . . . . . . . . . . . . . . . . . . . . 781 F. Rojo

4

Aerobic Degradation of Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . 799 D. Pe´rez-Pantoja . B. Gonza´lez . D. H. Pieper

5

Aerobic Degradation of Chloroaromatics . . . . . . . . . . . . . . . . . . . . . . . . . . 839 D. H. Pieper . B. Gonza´lez . B. Ca´mara . D. Pe´rez-Pantoja . W. Reineke

6

Aerobic Degradation of Halogenated Aliphatics . . . . . . . . . . . . . . . . . . . . . 865 S. Fetzner

Part 3 Biochemistry of Anaerobic Degradation . . . . . . . . . . . . . . . . . . . . . . . . . 887 7

The Biochemistry of Anaerobic Methane Oxidation . . . . . . . . . . . . . . . . . . 889 M. Taupp . L. Constan . S. J. Hallam

8

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes . . . . 909 F. Widdel . O. Grundmann

9

Anaerobic Degradation of Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . 925 M. Tierney . L. Y. Young

10

Microbial Degradation of Aliphatic and Aromatic Hydrocarbons with (Per)Chlorate as Electron Acceptor . . . . . . . . . . . . . . . . . . . . . . . . . . . 935 F. Mehboob . S. Weelink . F. T. Saia . H. Junca . A. J. M. Stams . G. Schraa

11

Hydrocarbon Degradation Coupled to Metal Reduction . . . . . . . . . . . . . . . 947 M. L. Heinnickel . F. M. Kaser . J. D. Coates

12

Anaerobic Degradation of Isoprene-Derived Compounds . . . . . . . . . . . . . . 957 J. Harder

13

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 963 D. Z. Sousa . M. Balk . M. Alves . B. Schink . M. J. McInerney . H. Smidt . C. M. Plugge . A. J. M. Stams

Table of Contents

Part 4 Enzymology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 981 14

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 983 F. Widdel . F. Musat

15

Anaerobic Degradation of Hydrocarbons: Mechanisms of C–H-Bond Activation in the Absence of Oxygen . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1011 M. Boll . J. Heider

16

The Role of Metals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1025 I. Bertini . A. Rosato

17

Biochemistry and Molecular Biology of Methane Monooxygenase . . . . . . 1045 J. C. Murrell . T. J. Smith

18

Aerobic Degradation of Aromatic Hydrocarbons: Enzyme Structures and Catalytic Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1057 J. D. Haddock

19

Oxidative Inactivation of Ring-Cleavage Extradiol Dioxygenases: Mechanism and Ferredoxin-Mediated Reactivation . . . . . . . . . . . . . . . . . 1071 Y. Jouanneau

20

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1081 J. Damborsky . R. Chaloupkova . M. Pavlova . E. Chovancova . J. Brezovsky

21

Lipolytic Enzymes from Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1099 S. Hausmann . K.-E. Jaeger

Part 5 Genetics (the Paradigms) (Section Editor: Victor de Lorenzo) . . . . . . . . . 1127 22

Transcriptional Control of the TOL Plasmid Pathways . . . . . . . . . . . . . . . 1129 P. Domı´nguez-Cuevas . S. Marque´s

23

Genetic Features and Regulation of n-Alkane Metabolism . . . . . . . . . . . . 1141 F. Rojo

24

Diversity of Naphthalene Biodegradation Systems in Soil Bacteria . . . . . 1155 A. M. Boronin . I. A. Kosheleva

25

Genomic View of Mycobacterial High Molecular Weight Polycyclic Aromatic Hydrocarbon Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1165 O. Kweon . S.-J. Kim . C. E. Cerniglia

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26

Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs . . . . . . 1179 M. Seeger . D. H. Pieper

27

Genetics and Molecular Features of Bacterial Dimethylsulfoniopropionate (DMSP) and Dimethylsulfide (DMS) Transformations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1201 J. M. Gonza´lez . A. W. B. Johnston . M. Vila-Costa . A. Buchan

28

Environmental Mining of Biological Activities on Hydrocarbons . . . . . . . . 1213 K. Watanabe

29

Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1219 R. R. Fulthorpe . E. M. Top

30

Experimental Evolution of Novel Regulatory Activities in Response to Hydrocarbons and Related Chemicals . . . . . . . . . . . . . . . . . . . . . . . . . 1235 V. Shingler

31

Rational Construction of Bacterial Strains with New/Improved Catabolic Capabilities for the Efficient Breakdown of Environmental Pollutants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1247 R.-M. Wittich . P. van Dillewijn . J.-L. Ramos

Part 6 Functional Genomics (the Paradigms) (Section Editor: Victor de Lorenzo) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1255 32

Bioinformatic, Molecular and Genetic Tools for Exploring Genome-wide Responses to Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . 1257 O. N. Reva . B. Tu¨mmler

33

Alcanivorax borkumensis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1265 V. Martins dos Santos . J. Sabirova . K. N. Timmis . M. M. Yakimov . P. N. Golyshin

34

Marinobacter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1289 R. Grimaud

35

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1297 J. I. Jime´nez . J. Nogales . J. L. Garcı´a . E. Dı´az

36

Genomics of Methylococcus capsulatus . . . . . . . . . . . . . . . . . . . . . . . . . . . 1327 J. C. Murrell

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37

Roseobacter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1335 A. Buchan . J. M. Gonza´lez

38

Rhodococcus: Genetics and Functional Genomics . . . . . . . . . . . . . . . . . . . 1345 M. J. Larkin . L. A. Kulakov . C. C. R. Allen

39

Phylogenomics of Aerobic Bacterial Degradation of Aromatics . . . . . . . . 1355 D. Pe´rez-Pantoja . R. Donoso . H. Junca . B. Gonza´lez . D. H. Pieper

40

Transcriptional Networks that Regulate Hydrocarbon Biodegradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1399 G. Carbajosa . I. Cases

41

Emerging Systems and Synthetic Biology Approaches to Hydrocarbon Biotechnology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1411 V. de Lorenzo . S. Fraile . J. I. Jime´nez

Part 7 Cellular Ecophysiology: Problems of Hydrophobicity, Bioavailability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1437 42

Introduction: Problems of Hydrophobicity/Bioavailability . . . . . . . . . . . . 1439 H. Harms . K. E. C. Smith . L. Y. Wick

43

Water-Hydrophobic Compound Interactions with the Microbial Cell . . . . 1451 E. M. McCammick . V. S. Gomase . T. J. McGenity . D. J. Timson . J. E. Hallsworth

44

Matrix-Hydrophobic Compound Interactions . . . . . . . . . . . . . . . . . . . . . . 1467 H. Harms . L. Y. Wick . K. E. C. Smith

45

Microorganism-Hydrophobic Compound Interactions . . . . . . . . . . . . . . . 1479 H. Harms . K. E. C. Smith . L. Y. Wick

46

Biofilm Development at Interfaces between Hydrophobic Organic Compounds and Water . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1491 R. Grimaud

47

Production and Roles of Biosurfactants and Bioemulsifiers in Accessing Hydrophobic Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1501 A. Perfumo . T. J. P. Smyth . R. Marchant . I. M. Banat

48

Uptake and Assimilation of Hydrophobic Substrates by the Oleaginous Yeast Yarrowia lipolytica . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1513 F. Thevenieau . A. Beopoulos . T. Desfougeres . J. Sabirova . K. Albertin . S. Zinjarde . J.-M. Nicaud

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Part 8 Cellular Ecophysiology: Uptake . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1529 49

Chemotaxis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1531 R. E. Parales . J. L. Ditty

50

Substrate Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1545 R. E. Parales . J. L. Ditty

51

Fungi as Transport Vectors for Contaminants and Contaminant-Degrading Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1555 L. Y. Wick . S. Furuno . H. Harms

Part 9 Cellular Ecophysiology: Problems of Solventogenicity, Solvent Tolerance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1563 52

Toxicity of Hydrocarbons to Microorganisms . . . . . . . . . . . . . . . . . . . . . . 1565 H. J. Heipieper . P. M. Martı´nez

53

Genetics of Accessing and Exploiting Hydrocarbons . . . . . . . . . . . . . . . . . 1575 C. Daniels . T. del Castillo . T. Krell . A. Segura . A. Busch . J. Lacal . J.-L. Ramos

54

Extrusion Pumps for Hydrocarbons: An Efficient Evolutionary Strategy to Confer Resistance to Hydrocarbons . . . . . . . . . . . . . . . . . . . . 1585 T. Krell . B. C ¸ ady´rcy´ . A. Segura . V. Garcı´a . C. Daniels . J.-L. Ramos

55

Membrane Composition and Modifications in Response to Aromatic Hydrocarbons in Gram Negative Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . 1595 A. Segura . P. Bernal . C. Pini . T. Krell . C. Daniels . J.-L. Ramos

56

Cis–Trans Isomerase of Unsaturated Fatty Acids: An Immediate Bacterial Adaptive Mechanism to Cope with Emerging Membrane Perturbation Caused by Toxic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . 1605 H. J. Heipieper . J. Fischer . F. Meinhardt

57

Surface Properties and Cellular Energetics of Bacteria in Response to the Presence of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1615 H. J. Heipieper . S. Cornelissen . M. Pepi

58

Microbiology of Oil Fly Larvae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1625 K. W. Nickerson . B. Plantz

Part 10 Cellular Ecophysiology: Problems of Feast or Famine . . . . . . . . . . . . . 1635 59

Kinetics and Physiology at Vanishingly Small Substrate Concentrations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1637 D. K. Button

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60

Feast: Choking on Acetyl-CoA, the Glyoxylate Shunt, and Acetyl-CoA-Driven Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1649 M. de la Pen˜a Mattozzi . Y. Kang . J. D. Keasling

61

Nitrogen Fixation and Hydrocarbon-Oxidizing Bacteria . . . . . . . . . . . . . . 1661 J. Foght

Volume 3 Microbes and Communities Utilizing Hydrocarbons, Oils and Lipids Part 1 The Microbes (Section Editor: Terry McGenity) . . . . . . . . . . . . . . . . . . . . 1669 1

Prokaryotic Hydrocarbon Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1671 R. C. Prince . A. Gramain . T. J. McGenity

2

Hydrocarbon-Degrading Sphingomonads: Sphingomonas, Sphingobium, Novosphingobium, and Sphingopyxis . . . . . . . . . . . . . . . . 1693 M. A. Kertesz . A. Kawasaki

3

Marine, Hydrocarbon-Degrading Alphaproteobacteria . . . . . . . . . . . . . . . 1707 S.-J. Kim . K. K. Kwon

4

Hydrocarbon Degradation by Betaproteobacteria . . . . . . . . . . . . . . . . . . 1715 R. E. Parales

5

Marinobacter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1725 R. Duran

6

Alcanivorax . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1737 S. Cappello . M. M. Yakimov

7

Oleiophilus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1749 S. Cappello . M. M. Yakimov

8

Oleispira . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1755 P. N. Golyshin . M. Ferrer . T. N. Chernikova . O. V. Golyshina . M. M. Yakimov

9

Thalassolituus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1765 M. M. Yakimov . M. Genovese . R. Denaro

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10

Neptunomonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1773 B. P. Hedlund . K. C. Costa

11

Cycloclasticus: A Genus of Marine Polycyclic Aromatic Hydrocarbon Degrading Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1781 J. T. Staley

12

Microbiology of Hydrocarbon-Degrading Pseudomonas . . . . . . . . . . . . . 1787 N. J. Palleroni . D. H. Pieper . E. R. B. Moore

13

Acinetobacter and Alkanindiges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1799 E. Ron . E. Rosenberg

14

Xanthomonads . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1805 H.-K. Chang . G. J. Zylstra

15

Bacteroidetes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1813 S.-J. Kim . K. K. Kwon

16

Actinobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1819 P. Ka¨mpfer

17

Rhodococcus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1839 M. J. Larkin . L. A. Kulakov . C. C. R. Allen

18

Micrococcineae: Arthrobacter and Relatives . . . . . . . . . . . . . . . . . . . . . . . 1853 C. T. Hennessee . Q. X. Li

19

Degradation of Polycyclic Aromatic Hydrocarbons by Mycobacterium Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1865 S.-J. Kim . O. Kweon . C. E. Cerniglia

20

Thermoleophilum: A Gram-Positive Hydrocarbonoclastic Thermophilic Bacterium . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1881 P. N. Golyshin . H. Lu¨nsdorf . M. Ferrer . M. M. Yakimov

21

The Genus Geobacillus and Hydrocarbon Utilization . . . . . . . . . . . . . . . . 1887 R. Marchant . I. M. Banat

22

Psychrophiles - Cold-Adapted Hydrocarbon-Degrading Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1897 A. Lo Giudice . V. Bruni . M. De Domenico . L. Michaud

23

Hydrocarbon-Degradation by Acidophilic Microorganisms . . . . . . . . . . . . 1923 W. F. M. Ro¨ling

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24

Alkaliphilic Hydrocarbon Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1931 T. J. McGenity . C. Whitby . A. Fahy

25

Halophilic Hydrocarbon Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1939 T. J. McGenity

26

The Aerobic Methane Oxidizing Bacteria (Methanotrophs) . . . . . . . . . . . . 1953 J. C. Murrell

27

Facultative Methane Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1967 S. N. Dedysh . P. F. Dunfield

28

Symbiotic Methane Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1977 J. M. Petersen . N. Dubilier

29

Anaerobic Hydrocarbon-Degrading Microorganisms: An Overview . . . . . 1997 F. Widdel . K. Knittel . A. Galushko

30

Anaerobic Methane Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2023 K. Knittel . A. Boetius

31

Nitrate, Perchlorate and Metal Respirers . . . . . . . . . . . . . . . . . . . . . . . . . 2033 F. M. Kaser . J. D. Coates

32

Anaerobic Utilization of Halohydrocarbons . . . . . . . . . . . . . . . . . . . . . . . 2049 S. H. Zinder

33

Eukaryotic Hydrocarbon Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2065 R. C. Prince

34

Degradation of Polycyclic Aromatic Hydrocarbons by Fungi . . . . . . . . . . 2079 C. E. Cerniglia . J. B. Sutherland

35

The Hydrocarbon-Degrading Oleaginous Yeast Yarrowia lipolytica . . . . . 2111 A. Beopoulos . T. Desfougeres . J. Sabirova . S. Zinjarde . C. Neuve´glise . J.-M. Nicaud

Part 2 Microbes Utilizing Non-Hydrocarbon Components of Fossil Fuels . . . . . 2123 36

Introduction to Microorganisms Utilizing Nitrogen and Sulfur Containing Heterocyclic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . 2125 S. Le Borgne

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37

Microorganisms Utilizing Sulfur-Containing Hydrocarbons . . . . . . . . . . . 2129 S. Le Borgne . M. Ayala

38

Microorganisms Utilizing Nitrogen-Containing Heterocyclic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2143 M. Morales . S. Le Borgne

Part 3 Microbial Communities Based on Hydrocarbons, Oils and Fats: Natural Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2159 39

Microbial Communities in Oil Shales, Biodegraded and Heavy Oil Reservoirs, and Bitumen Deposits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2161 J. Foght

40

Permafrost – Current and Future Challenges to Study Methanotrophy in Permafrost Affected Tundra and Wetlands . . . . . . . . . . . . . . . . . . . . . . 2173 S. Liebner . D. Wagner

41

Acidic Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2181 P. F. Dunfield . S. N. Dedysh

42

Habitats of Anaerobic Methane Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . 2193 A. Boetius . K. Knittel

43

Sulfate-Reducing and Methanogenic Hydrocarbon-Oxidizing Microbial Communities in the Marine Environment . . . . . . . . . . . . . . . . . 2203 A. Teske

44

Fungal Communities of Methane Clathrate-Bearing Deep Sea Sediments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2225 L. Cao

45

The Meta-Methanoxgenome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2231 M. Taupp . S. J. Hallam

46

Lipid Metabolism and the Rumen Microbial Ecosystem . . . . . . . . . . . . . . 2245 G. N. Jarvis . E. R. B. Moore

Part 4 Microbial Communities Based on Hydrocarbons, Oils and Fats: Anthropogenically-Created Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . 2259 47

The Oil Reservoir Ecosystem . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2261 B. Ollivier . D. Alazard

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48

Biodiesel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2271 R. C. Prince

49

Coal, Coal Mines and Spoil Heaps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2277 B. M. Kirby . C. J. Vengadajellum . S. G. Burton . D. A. Cowan

50

Microbial Hydrocarbon Degradation at Coal Gasification Plants . . . . . . . . 2293 R. U. Meckenstock . T. Lueders . C. Griebler . D. Selesi

51

Microbial Communities in Hydrocarbon-Contaminated Temperate, Tropical, Alpine, and Polar Soils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2313 C. W. Greer . L. G. Whyte . T. D. Niederberger

52

Bacterial Diversity in Hydrocarbon-Polluted Rivers, Estuaries and Sediments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2329 C. W. Greer

53

Impact of Pollution on Microbial Mats . . . . . . . . . . . . . . . . . . . . . . . . . . . 2339 R. Duran

54

Bacterial Communities in Hydrocarbon-Contaminated Marine Coastal Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2349 L. Berthe-Corti . M. Nachtkamp

55

Harbors and Marinas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2361 B. Nogales

56

The Microbiology of Metal Working Fluids . . . . . . . . . . . . . . . . . . . . . . . . 2369 I. P. Thompson . C. J. van der Gast

57

Milk Fat/Rancidity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2377 M. Jahn . D. Jahn

58

Vegetable Oil Wastes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2393 R. Denaro . S. Cappello . M. M. Yakimov

59

Foam in Wastewater Treatment Facilities . . . . . . . . . . . . . . . . . . . . . . . . . 2401 F. L. de los Reyes III

60

Functional Gene Diversity, Biogeography, Dynamics . . . . . . . . . . . . . . . . 2413 S. M. Nı´ Chadhain . G. J. Zylstra

61

Role of Protists in Microbial Interactions with Hydrocarbons . . . . . . . . . . 2423 T. Stoeck . V. Edgcomb

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Volume 4 Consequences of Microbial Interactions with Hydrocarbons, Oils and Lipids Part 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2435 1

Exploiting Microbial Diversity: The Challenges and the Means . . . . . . . . . 2437 V. de Lorenzo

Part 2 Applications: Organics Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . 2459 2

Bioremediation/Biomitigation: Introduction . . . . . . . . . . . . . . . . . . . . . . . 2461 E. Ron . E. Rosenberg

3

Reactive Tracers to Characterize Pollutant Distribution and Behavior in Aquifers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2465 R. D. Wilson

4

Natural Attenuation of Hydrocarbon Compounds in Groundwater . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2473 S. A. Banwart . S. F. Thornton

5

Weathered Hydrocarbon Biotransformation: Implications for Bioremediation, Analysis, and Risk Assessment . . . . . . . . . . . . . . . . . . . . 2487 K. J. Brassington . S. J. T. Pollard . F. Coulon

6

Role of Fertilizers: Biostimulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2501 E. Ron . E. Rosenberg

7

Cometabolic Bioremediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2505 T. C. Hazen

8

Role of Biosurfactants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2515 E. Ron . E. Rosenberg

9

Biostimulation Strategies for Enhanced Bioremediation of Marine Oil Spills Including Chronic Pollution . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2521 M. Nikolopoulou . N. Kalogerakis

10

Bioaugmentation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2531 N. Boon . W. Verstraete

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11

Plant-Microbe Partnerships . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2545 N. Weyens . S. Monchy . J. Vangronsveld . S. Taghavi . D. van der Lelie

12

Removal of Hydrocarbons and Other Related Chemicals via the Rhizosphere of Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2575 J. L. Ramos . E. Duque . P. van Dillewjin . C. Daniels . T. Krell . M. Espinosa-Urgel . M.-I. Ramos-Gonza´lez . S. Rodrı´guez . M. Matilla . R. Wittich . A. Segura

13

In Situ: Groundwater Bioremediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2583 T. C. Hazen

14

Remediation of Petrol and Diesel in Subsurface from Petrol Station Leaks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2597 R. C. Prince . G. S. Douglas

15

Remediation of BTEX in Groundwater Underlying Petrochemical Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2609 A. Fahy . T. J. McGenity

16

Bioremediation of Marine Oil Spills . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2617 R. C. Prince

17

Anaerobic Digesters for Digestion of Fat-Rich Materials . . . . . . . . . . . . . . 2631 M. Carballa . W. Vestraete

18

The Industrial Consequences of Microbial Deterioration of Metal-Working Fluid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2641 D. Theaker . I. Thompson

Part 3 Applications: Biomonitoring . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2651 19

Genetic Constructs: Molecular Tools for the Assembly of Environmental Bacterial Biosensors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2653 A. de las Heras . V. de Lorenzo

20

GeoChip: A High Throughput Genomic Tool for Linking Community Structure to Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2677 J. D. Van Nostrand . Y. Liang . Z. He . G. Li . J. Zhou

21

Q-RT-PCR Detection of Substrate-Specific Gene Expression . . . . . . . . . . . 2687 R. Denaro . M. M. Yakimov . M. Genovese

22

Antibody Microarrays for Environmental Monitoring . . . . . . . . . . . . . . . . 2699 V. Parro

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Part 4 Applications: Fuel Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2711 23

Using Microorganisms as Prospecting Agents in Oil and Gas Exploration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2713 C. Hubert . A. Judd

24

3 Oil Recovery: Fundamental Approaches and Principles of Microbially Enhanced Oil Recovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2727 H. Volk . P. Hendry

25

3 Oil Recovery: Experiences and Economics of Microbially Enhanced Oil Recovery (MEOR) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2739 H. Volk . K. Liu

26

Microbial Ecology of Oil Reservoir Souring and its Control by Nitrate Injection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2753 C. Hubert

27

Application of Microorganisms to the Processing and Upgrading of Crude Oil and Fractions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2767 M. Morales . M. Ayala . R. Vazquez-Duhalt . S. Le Borgne

28

Genetics Engineering for Removal of Sulfur and Nitrogen from Fuel Heterocycles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2787 E. Dı´az . J. L. Garcı´a

29

Biofuels (Butanol-Ethanol Production) . . . . . . . . . . . . . . . . . . . . . . . . . . . 2803 L. P. Wackett

30

Biomethane as an Energy Source . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2809 C. Bochiwal . C. O’Malley . J. P. J. Chong

31

Hydrocarbons from Algae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2817 J. G. Qin

32

Biodiesel from Microalgae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2827 A. Salis . M. Nicolo` . S. Guglielmino . V. Solinas

Part 5 Applications: Chemicals Production . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2841 33

Enzymatic Functionalization of Hydrocarbon-like Molecules . . . . . . . . . . 2843 N. Lo´pez-Corte´s . A. Beloqui . A. Ghazi . M. Ferrer

34

Screening for Enantioselective Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . 2859 B. Franken . K.-E. Jaeger . J. Pietruszka

Table of Contents

35

Biopetrochemicals via Biocatalysis by Hydrocarbons Microbes and their Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2877 K. Buehler . A. Schmid

36

Chemical Feedstocks and Fine Chemicals from Other Substrates . . . . . . . 2891 K. Muffler . N. Tippko¨tter . R. Ulber

37

Chemical Production – Biohalogenation . . . . . . . . . . . . . . . . . . . . . . . . . . 2903 C. D. Murphy . R. Grant

38

Metagenomic Mining of Enzyme Diversity . . . . . . . . . . . . . . . . . . . . . . . . 2911 M. E. Guazzaroni . A. Beloqui . J. M. Vieites . Y. Al-ramahi . N. L. Corte´s . A. Ghazi . P. N. Golyshin . M. Ferrer

39

Evolving Enzymes for Biocatalysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2929 U. T. Bornscheuer

40

Synthetic Biology for Biocatalysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2939 M. Bujara . S. Billerbeck . F. Greve . S. Panke

41

Microbial Production of Isoprenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2951 J. D. Keasling

42

Rediscovering Biopolymers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2967 S. A. Rivas . M. Bassas Galia`

43

Polyhydroxyalkanoates Produced by Hydrocarbon-Degrading Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2981 J. Sabirova

44

Biotechnological Production and Significance of Triacylglycerols and Wax Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2995 H. M. Alvarez

45

Yarrowia lipolytica as a Cell Factory for Oleochemical Biotechnology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3003 A. Beopoulos . T. Desfouge´res . J. Sabirova . J.-M. Nicaud

46

Ethylene Production by Fungi: Biological Questions and Future Developments Towards a Sustainable Polymers Industry . . . . . . . . . . . . . 3011 V. Chague´

47

Lipid-Containing Secondary Metabolites from Algae . . . . . . . . . . . . . . . . 3021 J. G. Qin

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48

Protein Emulsifiers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3031 E. Ron . E. Rosenberg

49

Rhamnolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3037 F. Leitermann . V. Walter . C. Syldatk . R. Hausmann

Part 6 Global Consequences of the Consumption and Production of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3053 50

Global Scale Consequences of Biological Methane Production . . . . . . . . . 3055 F. S. Colwell . W. Ussler III

51

Ecology of Aerobic Methanotrophs and their Role in Methane Cycling . . . 3067 Y. Chen . J. C. Murrell

52

Global Consequences of Anaerobic Methane Oxidation . . . . . . . . . . . . . . 3077 M. Strous

53

Global Consequences of the Microbial Production and Consumption of Inorganic and Organic Sulfur Compounds . . . . . . . . . . . . . . . . . . . . . . 3087 D. P. Kelly

54

Hydrocarbon Degradation in Petroleum Reservoirs . . . . . . . . . . . . . . . . . 3097 I. M. Head . S. R. Larter . N. D. Gray . A. Sherry . J. J. Adams . C. M. Aitken . D. M. Jones . A. K. Rowan . H. Huang . W. F. M. Ro¨ling

Part 7 Human-Animal-Plant Health and Physiology Consequences of Microbial Interactions with Hydrocarbons and Lipids . . . . . . . . . . . . . . 3111 55

Gastrointestinal Tract: Fat Metabolism in the Colon . . . . . . . . . . . . . . . . . 3113 L. Hoyles . R. J. Wallace

56

Gastrointestinal Tract: Intestinal Fatty Acid Metabolism and Implications for Health . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3119 L. Hoyles . R. J. Wallace

57

Gastrointestinal Tract: Microbial Metabolism of Steroids . . . . . . . . . . . . . 3133 P. Ge´rard

58

Obesity, Bacteria and Fat . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3141 C. Grootaert . T. Van de Wiele . W. Verstraete

Table of Contents

59

Conversion of Hydrocarbons by Gastrointestinal Microbiota and Consequences for Risk Assessment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3147 T. van de Wiele . W. Verstraete

60

Forage Lipids and Effects on Ruminant Productivity . . . . . . . . . . . . . . . . 3159 R. J. Dewhurst

61

Oral Microbiology: Pathogens, Methanogens, Sulfate-Reducing and Methylotrophic Bacteria in Halitosis and Periodontitis . . . . . . . . . . . . . . . 3167 A. P. Wood . D. P. Kelly

62

Lipid Rafts and Pseudomonas aeruginosa Infections . . . . . . . . . . . . . . . . . 3179 X. Li . Y. Zhang . E. Gulbins

63

Mycobacterial Lipid Bodies and the Chemosensitivity and Transmission of Tuberculosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3185 M. R. Barer . N. J. Garton

64

Lipids and Legionella Virulence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3195 O. Geiger

65

Skin Microbiology, Body Odor, and Methylotrophic Bacteria . . . . . . . . . . 3203 A. P. Wood . D. P. Kelly

66

Skin: Acne and Propionibacterium acnes Genomics . . . . . . . . . . . . . . . . . . 3215 H. Bru¨ggemann

67

Methylotrophic Bacteria in Trimethylaminuria and Bacterial Vaginosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3227 A. P. Wood . F. J. Warren . D. P. Kelly

68

Lipases as Pathogenicity Factors of Bacterial Pathogens of Humans . . . . 3241 J. Bender . A. Flieger

69

Lipases as Pathogenicity Factors of Fungi . . . . . . . . . . . . . . . . . . . . . . . . 3259 C. Gaillardin

70

Lipases as Pathogenicity Factors of Plant Pathogens . . . . . . . . . . . . . . . . 3269 S. Subramoni . Z. R. Sua´rez-Moreno . V. Venturi

71

Role of Cellular Control of Propionyl-CoA Levels for Microbial Pathogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3279 M. Brock

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72

Oil Degraders as Pathogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3293 F. Rojo . J. L. Martı´nez

73

Pathogens in Metal Working Fluids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3305 I. P. Thompson . C. J. van der Gast

74

Infection Prevention: Oil- and Lipid-Containing Products in Vaccinology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3311 T. Ebensen . B. Fuchs . K. Schulze . C. A. Guzma´n

Part 8 The Future . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3333 75

Biogas-Based Sustainable Bio-Economy . . . . . . . . . . . . . . . . . . . . . . . . . . 3335 W. Verstraete

76

Methane Production in a More Saline World . . . . . . . . . . . . . . . . . . . . . . 3337 T. J. McGenity

77

Potential for Microbial Interventions to Reduce Global Warming . . . . . . . 3339 D. P. Kelly . A. P. Wood

78

Can we Improve Bioremediation? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3351 R. C. Prince

79

Role of Plant-Microbe Partnerships to Deal with Environmental Challenges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3357 N. Weyens . S. Monchy . J. Vangronsveld . S. Taghavi . D. van der Lelie

80

Natural Biotechnology: Exploiting Microbial Diversity . . . . . . . . . . . . . . . 3361 A. J. M. Stams

81

‘‘Sequence First, Ask Questions Later’’: The Impact of Next Generation-Omics on the Discovery of Novel Microbial and Lipid Hydrocarbon Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3367 D. A. Cowan . I. M. Tuffin . F. T. Robb

82

Biopolymers from Sunshine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3375 S. Arias Rivas . M. Bassas Galia`

83

Neutral Lipid Metabolism in Yeast as a Template for Biomedical Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3381 K. Athenstaedt

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84

Research Needs in Vaccinology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3383 Carlos A. Guzma´n

85

Personal Care Products . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3387 A. P. Wood

Volume 5 Experimental Protocols and Appendices Part 1 Study Systems (Section Editor: Jan Roelof van der Meer) . . . . . . . . . . . . 3395 1

Field Studies – Demonstrating the Efficacy of Bioremediation . . . . . . . . . 3397 R. C. Prince

2

Groundwater Sampling for Nucleic Acid Biomarker Analysis . . . . . . . . . . 3407 K. M. Ritalahti . J. K. Hatt . E. Petrovskis . F. E. Lo¨ffler

3

Deep Sea Sampling, Sample Work-up and Analysis . . . . . . . . . . . . . . . . . 3419 S. Borin . D. Daffonchio

4

Sampling the Deep Sub-Surface Using Drilling and Coring Techniques . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3427 T. L. Kieft

5

Methods for the Study of Cold Seep Ecosystems . . . . . . . . . . . . . . . . . . . 3443 A. Boetius . F. Wenzho¨fer

6

Microcosm Experiments for Simulation of Freeze-Thaw Cycles and Studying Methane Dynamics in Permafrost-Affected Soils . . . . . . . . . . . . 3453 D. Wagner

7

Combined Use of Fluorescent Reporters and Flow Cytometry for Simultaneous Monitoring of Bacterial Growth and Gene Expression on Plant Roots . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3461 L. Rochat . M. Pe´chy-Tarr . M. Maurhofer . C. Keel

8

Accidental and Deliberate Oil Spills in Europe: Detection, Sampling and Subsequent Analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3471 L. Peperzak . P. Kienhuis . C. P. D. Brussaard . J. Huisman

9

An Experimental Oil Spill at Sea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3491 C. P. D. Brussaard . L. Peperzak . Y. Witte . J. Huisman

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10

In Situ Microcosm Studies to Characterize Microbial Processes in the Field . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3503 M. Ka¨stner . H. H. Richnow

11

Mesocosms for Oil Spill Simulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3513 S. Cappello . M. M. Yakimov

12

Microcosms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3523 A. Fahy . B. McKew

13

Growth of Hydrocarbon-Degrading Bacteria in Continuous Culture . . . . . 3529 M. Bucheli-Witschel . T. Egli

14

Microcosms for Biofilm Analysis on Hydrophobic Substrates – A Multiple Approach to Study Biodiversity, Metabolic Activity and Biofilm Structure and Dynamic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3543 A. J. Macedo . W.-R. Abraham

15

‘‘Clay Hutches,’’ a Soil Derived Primordial Bacterium-Mineral Interaction – an Ultrastructural Approach . . . . . . . . . . . . . . . . . . . . . . . . 3553 H. Lu¨nsdorf

16

Microcolony Growth Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3561 F. Reinhard . J. R. van der Meer

Part 2 Analytical Procedures (Section Editor: Jan Roelof van der Meer) . . . . . . . 3573 17

Chemical Analysis of Hydrocarbons in Petroleum Oils and the Assessment of Environmental Contamination . . . . . . . . . . . . . . . . . . . . . 3575 J. W. Readman

18

Solid Phase Microextraction (SPME) for Determining the Freely Dissolved Concentrations of Oil Hydrocarbons . . . . . . . . . . . . . . . . . . . . . 3583 K. E. C. Smith . L. Y. Wick

19

Measuring Hydrocarbons in the Atmosphere . . . . . . . . . . . . . . . . . . . . . . 3593 I. Colbeck

20

Natural Stable Isotope Fractionation for the Assessment of Hydrocarbon Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3603 R. U. Meckenstock . H. H. Richnow

21

Respiration Rate Determination by Phosphorescence-Based Sensors . . . . 3613 T. J. Strovas . M. E. Lidstrom

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22

Determination of Aerobic Degradation Rates and Kinetics of Gaseous Hydrocarbons in Soil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3623 P. Ho¨hener

23

Determining the Tendency of Microorganisms to Interact with Hydrocarbon Phases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3631 H. Harms . L. Y. Wick

24

Bacterial Bioreporter Assays to Measure Hydrocarbons . . . . . . . . . . . . . . 3641 R. Tecon . S. Beggah . J. R. van der Meer

25

Microsensor Techniques to Study in situ Bacterial Metabolic Processes in Hydrocarbon-Polluted Marine Cyanobacterial Mats . . . . . . . . . . . . . . . 3655 R. M. M. Abed . D. de Beer

26

RNA Extraction and cDNA Analysis for Quantitative Assessment of Biomarker Transcripts in Groundwater . . . . . . . . . . . . . . . . . . . . . . . . . . . 3671 K. M. Ritalahti . C. Cruz-Garcı´a . E. Padilla-Crespo . J. K. Hatt . F. E. Lo¨ffler

27

Isolation and Analysis of Lipopeptides and High Molecular Weight Biosurfactants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3687 T. J. P. Smyth . A. Perfumo . S. McClean . R. Marchant . I. M. Banat

28

Isolation and Analysis of Low Molecular Weight Microbial Glycolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3705 T. J. P. Smyth . A. Perfumo . R. Marchant . I. M. Banat

29

Isolation and Analysis of Storage Compounds . . . . . . . . . . . . . . . . . . . . . 3725 M. Bassas Galia`

30

Isolation and Analysis of Lipids, Biomarkers . . . . . . . . . . . . . . . . . . . . . . . 3743 H. J. Heipieper

31

The Evaporative Weathering of Oil . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3751 K. E. C. Smith

32

Geological and Hydrogeological Characterization of Subsurface . . . . . . . 3759 D. Hunkeler

Part 3 Microbiology and Community Procedures (Section Editor: Jan Roelof van der Meer) . . . . . . . . . . . . . . . . . . . . . . . . 3769 33

Biodegradation Experiments – Classical Set-Up: Isolation of Aerobic, Xenobiotic-Degrading Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . 3771 B. Geueke . H.-P. E. Kohler

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34

Enrichment and Isolation of Hydrocarbon Degraders . . . . . . . . . . . . . . . . 3777 Z. Shao

35

Cultivation of Anaerobic Microorganisms with Hydrocarbons as Growth Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3787 F. Widdel

36

Methods for Studying Methanogens and Methanogenesis in Marine Sediments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3799 R. John Parkes . H. Sass . G. Webster . A. J. Watkins . A. J. Weightman . L. A. O’Sullivan . B. A. Cragg

37

Isolation and Characterization of Methanotrophs and Methylotrophs: Diversity of Methylotrophic Organisms and of One-Carbon Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3827 D. P. Kelly . A. P. Wood

38

Cultivation of Halophilic Hydrocarbon Degraders . . . . . . . . . . . . . . . . . . 3847 T. J. McGenity . A. Gramain

39

Enrichment and Isolation of Metal Respirers and Hydrocarbon Oxidizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3855 F. M. Kaser . J. D. Coates

40

Sampling, Isolation, Cultivation, and Characterization of Piezophilic Microbes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3869 S.-J. Kim . C. Kato

41

Studying the In Situ Role of Protistan Communities in Hydrocarbon Contaminated Water Samples via Community Profiling and CARD-FISH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3883 A. Behnke . M. Engel . V. Edgcomb . T. Stoeck

42

Microbial Oil Degradation Under Methanogenic Conditions . . . . . . . . . . . 3905 A. Sherry . N. Gray . C. Aitken . J. Dolfing

43

Two-Phase Cultivation Techniques for Hydrocarbon-Degrading Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3919 L. Y. Wick . C. Holliger

44

Ecophysiological Characterization of Substrate Affinities of Hydrocarbon Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3927 D. K. Button

45

Chemotactic Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3933 R. K. Jain . J. Pandey

Table of Contents

46

Protocols for the Characterization of Solvent Tolerant Microorganisms: Construction and Characterization of Mutants . . . . . . . . 3957 E. Duque . J. de la Torre . V. Garcı´a . C. Pini . S. Rodrı´guez-Conde . P. Godoy . M. A. Henares-Molina . T. Krell . C. Daniels . J. L. Ramos . A. Segura

47

Clone Libraries of Ribosomal RNA Gene Sequences for Characterization of Bacterial and Fungal Communities . . . . . . . . . . . . . . 3969 M. B. Leigh . L. Taylor . J. D. Neufeld

48

Real-Time PCR Approaches for Analysis of Hydrocarbon-Degrading Bacterial Communities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3995 B. A. McKew . C. J. Smith

49

Stable Isotope Probing of Hydrocarbon-Degraders . . . . . . . . . . . . . . . . . 4011 T. Lueders

50

Raman FISH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4027 D. S. Read . W. E. Huang . A. S. Whiteley

51

GeoChips for Analysis of Microbial Functional Communities . . . . . . . . . . 4039 J. D. Van Nostrand . L. Wu . Z. He . J. Zhou

52

Assessing Functionality by Differential Display and RNA Arbitrary PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4051 S. Bordenave . M. Gon˜i-Urriza . R. Duran

53

Biomonitoring by Antibody Microarrays . . . . . . . . . . . . . . . . . . . . . . . . . . 4063 V. Parro

54

Examination of Microbial Communities on Hydrocarbons by Means of Laser Scanning Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4073 T. R. Neu . J. R. Lawrence

55

Catalyzed Reporter Deposition- Fluorescence In Situ Hybridization (CARD-FISH) and Abundance of Cycloclasticus . . . . . . . . . . . . . . . . . . . . . 4085 E. Teira

56

Combined Microautoradiography and Fluorescence in situ Hybridization (MAR-FISH) for the Identification of Metabolically Active Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4093 J. L. Nielsen . P. H. Nielsen

57

Functional Flow Cytometry in Environmental Microbiology . . . . . . . . . . . 4103 S. Mu¨ller

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58

Molecular Profiling of Bacterial Communities via 16S rRNA Gene Based Approaches – Focus T-RFLP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4113 S. Paı¨sse´ . M. S. Gon˜i-Urriza . A. Fahy . R. Duran

59

Identification of Environmental Microorganisms by Fluorescence in situ Hybridization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4127 A. Pernthaler

60

Denaturing Gradient Gel Electrophoresis (DGGE) for Microbial Community Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4137 S. J. Green . M. B. Leigh . J. D. Neufeld

61

Introduction to Microplate MPN-Enumeration of Hydrocarbon Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4159 A. R. Johnsen

Part 4 Biochemical Procedures (Section Editor: Jan Roelof van der Meer) . . . . . 4173 62

University of Minnesota Biocatalysis/Biodegradation Database (UM-BBD) and Hydrocarbon Research . . . . . . . . . . . . . . . . . . . . . . . . . . . 4175 L. P. Wackett . L. B. M. Ellis

63

Procedures for Protein Isolation in Pure Culture and Microbial Communities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4183 A. Beloqui . M. E. Guazzaroni . M. Ferrer

64

Enzyme Assays, Substrate Specificities, Kinetic Parameters: Measurement of Enzyme Activities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4195 B. Geueke . H.-P. E. Kohler

65

Bacterial Solvent Responses and Tolerance: Cis–Trans Isomerization . . . . 4203 H. J. Heipieper . J. Fischer

66

Measurement of Hydrocarbon Transport in Bacteria . . . . . . . . . . . . . . . . 4213 J. L. Ditty . N. N. Nichols . R. E. Parales

67

Isolation and Characterization of Lipid Particles from Yeast . . . . . . . . . . . 4223 K. Athenstaedt

68

Microcalorimetry as a General Technique to Characterize Ligand Binding: What Needs to be Considered When Analyzing Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4231 T. Krell . M.-E. Guazzaroni . A. Busch . J. Lacal . W. Tera´n . S. Fillet . H. Silva-Jime´nez . J. L. Ramos

Table of Contents

Part 5 Genetic and System Procedures (Section Editor: Jan Roelof van der Meer) . . . . . . . . . . . . . . . . . . . . . . . . 4243 69

Genetic Analysis of Gram-Negative Bacteria Using Mini Tn5 Transposons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4245 A. Cebolla . M. Are´valo-Rodrı´guez

70

Multiple Displacement Amplification . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4255 J. A. Gilbert . K. Zhang . J. D. Neufeld

71

Genomics – Bacterial Genome Sequencing and Annotation . . . . . . . . . . . 4265 S. Schneiker-Bekel . T. Bekel . A. Pu¨hler

72

Genome Annotation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4281 B. Tu¨mmler

73

Functional Marker Gene Assays for Hydrocarbon Degrading Microbial Communities: Aerobic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4289 H. Junca . D. H. Pieper

74

Tools for Comparison of Bacterial Genomes . . . . . . . . . . . . . . . . . . . . . . . 4313 T. M. Wassenaar . T. T. Binnewies . P. F. Hallin . D. W. Ussery

75

Genome-Scale Constraint-Based Models to Navigate the Microbial Landscape . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4329 J. Puchałka . C. M. C. Lam . V. A. P. Martins dos Santos

76

Flux-Related Metabolic Network Descriptions . . . . . . . . . . . . . . . . . . . . . 4339 W. F. M. Ro¨ling

77

Aquatic Metagenome Library (Archive; Expression) Generation and Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4347 J. Gilbert

78

Transcriptome Analysis Using High-Density Oligonucleotide Microarrays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4353 D. R. Johnson

79

Use of Microarrays to Study Bacterial Responses to Hydrocarbons . . . . . 4367 G. Navarro-Avile´s . J. J. Rodrı´guez-Herva . J. Luis Ramos

80

A Methodology Applying Two-Dimensional Gel Electrophoresis to Analyze Bacterial Response to Contact with Hydrocarbon-Water Interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4377 P. J. Vaysse . R. Grimaud

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81

Proteogenomics to Study the Anaerobic Degradation of Aromatic Compounds and Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4385 R. Rabus . K. Trautwein

82

Interactomics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4407 T. Dammeyer . M. Schobert

83

Random and Site-Directed Mutagenesis of Transcriptional Regulator Proteins Implicated in Hydrocarbon Degradation Pathways . . . . . . . . . . . 4429 C. Vogne . S. Beggah . J. van der Meer

Part 6 Application Procedures (Section Editor: Jan Roelof van der Meer) . . . . . 4445 84

Commercial Application of Bioluminescence Full Cell Bioreporters for Environmental Diagnostics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4447 E. E. Diplock . H. A. Alhadrami . G. I. Paton

85

Human Cell Line Based Assays for Toxic Poly Halogenated and Poly Aromatic Hydrocarbons in Oils and Lipids . . . . . . . . . . . . . . . . . . . . . . . . 4459 B. van der Burg . A. Jonas . E. Sonneveld

86

Ecological Risk Assessment: The Triad Approach . . . . . . . . . . . . . . . . . . . 4465 M. Wagelmans

87

Assessment of Genotoxicity Following Exposure to Hydrocarbons: The Micronucleus Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4473 T. Galloway . C. Lewis . J. Hagger

88

Zebrafish Embryo Toxicity Assay, Combining Molecular and Integrative Endpoints at Various Developmental Stages . . . . . . . . . . . . . . . . . . . . . . 4481 J. R. K. Njiwa . M. J.-F. Suter . R. I. Eggen

89

Petroleum Toxicity and Bioaccumulation Studies in Fish, Sea Urchins and Mussels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4491 S. Cappello . M. M. Yakimov

90

Phytoplankton Viability Assay for Oil Compounds in Water . . . . . . . . . . . 4499 L. Peperzak . C. P. D. Brussaard

91

Intrinsic Bioremediation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . 4509 J. B. M. van Bemmel

92

Biostimulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4517 T. C. Hazen

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Bioaugmentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4531 M. L. B. da Silva . P. J. J. Alvarez

94

Bioremediation: Slow-Release Inoculation by Hot Spot Tubes . . . . . . . . . 4545 N. Boon

95

Isolation and Characterization of Microorganisms for Fuel Quality Improvement . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4553 M. Morales . S. Le Borgne

96

Genetic Traps for Surveying New Catalysts in (Meta) Genomic DNA . . . . . 4563 C. A. Carren˜o . V. de Lorenzo

97

Protocols to Screen for Enantioselective Lipases . . . . . . . . . . . . . . . . . . . 4581 B. Franken . K.-E. Jaeger . J. Pietruszka

98

Bacterial Secretion Systems for Use in Biotechnology: Autotransporter-Based Ultra-High Throughput Cell-Surface Display and Screening of Large Protein Libraries . . . . . . . . . . . . . . . . . . . . . . . . . 4587 S. Wilhelm . H. Kolmar . F. Rosenau

99

Engineering Monomer Composition of PHA Accumulated in Yeast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4601 C. W. J. McChalicher . F. Srienc

Part 7 Appendices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4609 Subject Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4637

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List of Contributors

Raeid M. M. Abed Biology Department, College of Science Sultan Qaboos University 123 Al Khoud Sultanate of Oman [email protected] Wolf-Rainer Abraham Helmholtz Centre for Infection Research Chemical Microbiology Inhoffenstrasse 7 38124 Braunschweig Germany [email protected] Jennifer J. Adams Petroleum Reservoir Group Department of Geology and Geophysics and Alberta Ingenuity Center for In situ Energy University of Calgary Calgary, AB Canada Carolyn M. Aitken School of Civil Engineering and Geosciences Newcastle University Newcastle upon Tyne NEI 7RU UK Dider Alazard Laboratoire de Microbiologie IRD, UMR 180, Universite´s de Provence et de la Me´diterrane´e ESIL Case 925 163 Avenue de Luminy

13288 Marseille cedex 9 France Koos Albertin Department of Microbial, Biochemical and Food Biotechnology University of the Free State Bloemfontein, 9300 South Africa H. A. Alhadrami Institute of Biological and Environmental Sciences Cruickshank Building University of Aberdeen Aberdeen, AB24 3UU UK Djamila Al-Halbouni Institute for Biology, Unit of Soil Ecology RWTH Aachen University Worringer Weg 1 52056 Aachen Germany Christopher C. R. Allen School of Biological Sciences and The QUESTOR Centre The Queen’s University of Belfast Belfast BT9 5AG, Northern Ireland UK Yamal Al-ramahi CSIC, Institute of Catalysis 28049 Madrid Spain

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List of Contributors

Hector M. Alvarez Regional Center for Research and Development (CRIDECIT) Faculty of Natural Science University of Patagonia San Juan Bosco 9000 Comodoro Rivadavia, Chutbut Argentina [email protected] Pedro J. J. Alvarez Department of Civil and Environmental Engineering Rice University Houston, TX 77005 USA [email protected] M. Alves Institute for Biotechnology and Bioengineering Centre of Biological Engineering University of Minho Braga Portugal Gregor Anderluh Department of Biology University of Ljubljana Veena pot 111 1000 Ljubljana Slovenia [email protected] Miguel Are´valo-Rodrı´guez Biomedal S. L. 41092 Seville Spain Karin Athenstaedt Institute of Biochemistry University of Technology Graz Petersgasse 12/2 8010 Graz Austria [email protected]

Marcela Ayala Department of Cellular Engineering and Biocatalysis Instituto de Biotecnologia, UNAM Cuernavaca Mexico Melike Balk Laboratory of Microbiology Wageningen University Dreijenplein 10 6703 HB Wageningen The Netherlands Ibrahim M. Banat School of Biomedical Sciences University of Ulster Coleraine, County Londonderry BT52 1SA Northern Ireland UK [email protected] Steven A. Banwart Groundwater Protection and Restoration Group Kroto Research Institute The University of Sheffield Sheffield S37HQ UK [email protected] Michael R. Barer Department of Infection, Immunity and Inflammation University of Leicester Medical School Maurice Shock Building University Road Leicester, LE19HN UK and Department of Clinical Microbiology University Hospitals of Leicester NHS Trust Leicester UK [email protected]

List of Contributors

Mo´nica Bassas Galia` Environmental Microbiology Laboratory Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany [email protected] Siham Beggah Department of Fundamental Microbiology, University of Lausanne Baˆtiment Biophore, Quartier UNIL-Sorge 1015 Lausanne Switzerland Anke Behnke Department of Biology University of Kaiserslautern 67663 Kaiserslautern Germany [email protected] Thomas Bekel Center for Biotechnology (CeBiTec) Bielefeld University 33594 Bielefeld Germany Ana Beloqui Department of Applied Biocatalysis CSIC, Institute of Catalysis 28049 Madrid Spain James Bender Robert Koch-Institute Berlin Germany Alhanasios Beopoulos Laboratoire de Microbiologie et Ge´ne´tique Mole´culaire INRA, UMR 1285, CNRS, UMR 2585 AgroParisTech Centre de Biotechnologie AgroIndustrielle 78850 Thiverval-Grignon France

Patricia Bernal Department of Environmental Protection CSIC Granada Spain Luise Berthe-Corti Institute for Chemistry and Biology of the Marine Environment (ICBM) University of Oldenburg Carl-von-Ossietzky-Str. 9-11 26129 Oldenburg Germany [email protected] Ivano Bertini Magnetic Resonance Center (CERM) University of Florence Sesto Fiorentino, Italy and Department of Chemistry University of Florence Via Luigi Sacconi 6 Sesto Fiorentino Italy [email protected] Sonja Billerbeck Bioprocess Laboratory Institute of Process Engineering ETH Zurich, Zurich Switzerland [email protected] Tim T. Binnewies Center for Biological Sequence Analysis Technical University of Denmark Lyngby 2800 Denmark and Roche Diagnostics Ltd. Advanced Systems Group, Global Platforms & Support Forrenstrasse 6343 Rotkreuz Switzerland

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Russel E. Bishop Department of Biochemistry and Biomedical Sciences McMaster University 4 H19 Health Science Centre 1200 Main Street West Hamilton, ON LBN 3Z5 Canada [email protected] Chitran Bochiwal Department of Biology University of York York YO 10 5YW UK Antje Boetius Max Planck Institute for Marine Microbiology Celsiusstr. 1 28359 Bremen Germany and Alfred Wegener Institute for Marine and Polar Research Bremerhaven Germany [email protected]

Sylvain Bordenave Equipe Environnement et Microbiologie UMR CNRS IPREM 5254 Universite´ de Pau et des Pays de l’Adour IBEAS BP1155 Pau cedex France [email protected] Sara Borin Department of Food Science and Microbiology University of Milan Via Celoria 2 20133 Milan Italy [email protected] Uwe T. Bornscheuer Department of Biotechnology and Enzyme Catalysis Institute of Biochemistry Greifswald University Felix-Hausdorff-Str.4 17487 Greifswald Germany [email protected]

Matthias Boll Institute of Biochemistry University of Leipzig 04103 Leipzig Germany [email protected]

Alexander M. Boronin Skryabin Institute of Biochemistry and Physiology of Microorganisms Russian Academy of Sciences Pushchino State University 142290 Pushchino Moscow Russia [email protected]

Nico Boon Laboratory of Microbial Ecology and Technology (LabMET) Ghent University Coupure L653 9000 Ghent Belgium [email protected]

Kirsty J. Brassington Centre for Resource Management and Efficiency Sustainable Systems Department School of Applied Sciences Cranfield University Cranfield MK 43 OAL UK

List of Contributors

Jan Brezovsky Loschmidt Laboratories Institute of Experimental Biology and National Centre for Biomolecular Research Masaryk University Kamenice 5/A4 625 00 Brno Czech Republic Matthias Brock Leibniz Institute for Natural Product Research and Infection Biology Hans Knoell Institute Beutenbergstr. 11a 07745 Jena Germany [email protected] Holger Bru¨ggemann Department of Molecular Biology Max Planck Institute for Infection Biology Charite´platz 1 10117 Berlin Germany [email protected] Andreas Brune Department of Biogeochemistry Max Planck Institute for Terrestrial Microbiology Karl-von-Frisch-Straße 35043 Marburg Germany [email protected] Vivia Bruni Department of Animal Biology and Marine Ecology University of Messina Messina Italy Corina P. D. Brussaard Royal Netherlands Institute for Sea Research/NIOZ Landsdiep 4 1790 AB Den Burg

Texel The Netherlands [email protected] Alison Buchan Department of Microbiology M409 Walters Life Sciences University of Tennessee Knoxville 37996, TN USA [email protected] Margarette Bucheli-Witschel Department of Environmental Microbiology Eawag, Swiss Federal Institute of Aquatic Science and Technology U¨berlandstrasse 133 8600 Dubendorf Switzerland Katja Buehler Department of Chemical Biotechnology Laboratory of Chemical and Biochemical Engineering Dortmund University of Technology Emil-Figge-Strasse 66 44227 Dortmund Germany Matthias Bujara Bioprocess Laboratory Institute of Process Engineering Eidgeno¨ssische Technische Hochschule (ETH) Zurich Zurich Switzerland [email protected] Stephanie G. Burton Department of Chemical Engineering Biocatalysis and Technical Biology Unit University of Cape Town, Rondebosch 7700, Cape Town South Africa

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Andreas Busch Department of Environmental Protection Estacioˇn Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas 18008 Granada Spain Don K. Button Institute of Marine Science University of Alaska Fairbanks, AK USA [email protected]

Marta Carballa Laboratory of Microbial Ecology and Technology (LabMET) Ghent University Ceupure Links 653 9000 Ghent Belgium [email protected] Carlos A. Carrenˇo Systems Biology Program Centro Nacional de Biotecnologı´a-CSIC 28049 Madrid Spain

Bilge C¸ady´rcy´ Department of Environmental Protection CSIC Granada Spain

Ildefonso Cases Spanish National Cancer Research Centre C/Melchor Ferna´ndez Almagro, 3 28029 Madrid Spain [email protected]

Beatriz Ca´mara Chemical Microbiology Bergische Universita¨t Wuppertal Gaußstr. 20 42097 Wuppertal Germany

Teresa del Castillo Department of Environmental Protection CSIC Granada Spain

Lixiang Cao Department of Biochemistry School of Life Sciences Zhongshan (Sun Yatsen) University Guangzhou China [email protected] Simone Cappello Istituto per l’Ambiente Marino Costiero (IAMC) of Messina IST-CNR Spianata S. Raineri 86 98122 Messina Italy [email protected] Guillernio Carbajosa Spanish National Cancer Research Centre C/Melchor Ferna´ndez Almagro, 3 28029 Madrid Spain [email protected]

Jean-Luc Cayol Laboratoire de Microbiologie IRD, UMR 180 Universite´s de Provence et de la Me´diterrane´e ESIL 13288 Marseille Cedex 9 France Angel Cebolla Biomedal, SL Avda. Ame´rico Vespucio 41092 Sevilla Spain [email protected] Carl E. Cerniglia Division of Microbiology National Center for Toxicological Research Food and Drug Administration 3900 NCTR Road, HFT-250 Jefferson, AR USA [email protected]

List of Contributors

Veronique Chague´ 50 bis chemin de la Garenne 91290 La Norville France [email protected] Radka Chaloupkova Loschmidt Laboratories Institute of Experimental Biology and National Centre for Bimolecular Research Masaryk University Kamenice 5/A4 625 00 Brno Czech Republic Hung-Kuang Chang Biotechnology Center for Agriculture and the Environment Rutgers University New Brunswick, NJ USA Y. Chen Department of Biological Sciences University of Warwick Coventry UK Tatyana N. Chernikova Environmental Microbiology Laboratory HZI-Helmholtz Centre for Infection Research 38124 Braunschweig Germany [email protected] James P. J. Chong Department of Biology University of York YO 10 5YW York UK [email protected] Eva Chovancova Loschmidt Laboratories Institute of Experimental Biology and National Centre for Biomolecular Research Masaryk University Kamenice 5/A4 625 00 Brno Czech Republic

John D. Coates Department of Plant and Microbial Biology University of California 271 Koshland Hall Berkeley, CA USA [email protected]

Ian Colbeck Department of Biological Sciences University of Essex Central Campus Wivenhole Park Colchester UK [email protected]

Frederick S. Colwell College of Oceanic and Atmospheric Sciences Oregon State University 104 COAS, Admin Bldg. Corvallis, OR USA [email protected]

Lea Constan Department of Microbiology and Immunology University of British Columbia Life Sciences Centre 2552-2350 Health Sciences Mall Vancouver, BC V6T 1Z3 Canada

Sjef Cornelissen Laboratory of Chemical Biotechnology Faculty of Biochemical and Chemical Engineering Technical University Dortmund Emil Figge Str. 66 44221 Dortmund Germany

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Nieves L. Corte´s CSIC, Institute of Catalysis Department of Applied Biocatalysis Marie Curie 2 28049 Madrid Spain Kyle C. Costa Department of Microbiology University of Washington Seattle WA, 98195-7242 USA [email protected] Fre´de´ric Coulon Centre for Resource Management and Efficiency Sustainable Systems Department, School of Applied Sciences Cranfield University Cranfield UK [email protected] Don A. Cowan Department of Biotechnology Institute for Microbial Biotechnology and Metagenomics University of the Western Cape Bellville, Cape Town South Africa [email protected]

Marcio L. B. da Silva Department of Civil and Environmental Engineering Rice University Houston, TX 77005 USA [email protected] Daniele Daffonchio Department of Food Science and Microbiology University of Milan Via Celoria 2 Milan Italy [email protected] Jiri Damborsky Loschmidt Laboratories Institute of Experimental Biology and National Centre for Biomolecular Research Masaryk University Kamenice 5/A4 625 00 Brno Czech Republic [email protected] Thorben Dammeyer Environmental Microbiology Laboratory Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany

Barry A. Cragg School of Earth and Ocean Sciences Cardiff University Main Building Park Place Cardiff, Wales CF10 3YE UK

Craig Daniels Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain

Claribel Cruz-Garcı´a School of Civil and Environmental Engineering Georgia Institute of Technology 311 Ferst Drive Atlanta, GA 30332 USA

Dirk de Beer Max-Planck-Institute for Marine Microbiology Celsiusstraße 1 28359 Bremen Germany [email protected]

List of Contributors

Maria De Domenico Department of Animal Biology and Marine Ecology University of Messina Messina Italy M. de la Penˇa Mattozzi Department of Plant and Microbial Biology University of California Berkeley CA 94720 USA Aitor de las Heras Systems Biology Program Centro Nacional de Biotecnologı´a-CSIC Campus de Cantoblanco Madrid 28049 Spain Vı´ctor de Lorenzo Systems Biology Program Centro Nacional de Biotecnologı´a-CSIC Campus de Cantoblanco Madrid 28049 Spain [email protected] Francis L. de los Reyes III Civil, Construction, and Environmental Engineering North Carolina State University 319B Mann Hall Raleigh, NC 27695 USA [email protected] Diego de Mendoza Instituto de Biologı´a Molecular y Celular de Rosario (IBR-CONICET) Departamento de Microbiologı´a, Facultad de Ciencias Bioquı´micas y Farmace´uticas Universidad Nacional de Rosario Suipacha, Rosario Argentina [email protected]

Svretlana N. Dedysh S.N. Winogradsky Institute of Microbiology Russian Academy of Sciences Prospect 60-Letya Octyabrya 7/2 Moscow 117312 Russia [email protected] Renata Denaro Institute for Coastal Marine Environment CNR (National Research Council) Department of Earth and Environment Raineri 86 98122 Messina Italy [email protected] Thomas Desfougeres Laboratoire de Microbiologie et Ge´ne´tique Mole´culaire, INRA AgroParisTech, Centre de Biotechnologie Agro-Industrielle INRA,UMR 1238, CNRS, UMR 2585 BP 01 78850 Thiverval-Grignon France Richard J. Dewhurst Animal Bioscience Centre Teagasc Dunsany, County Meath Ireland [email protected] Eduardo Diaz Department of Molecular Microbiology Centro de Investigationes Biolo´gicas Consejo Superior de Investigaciones Cientı´gicas Madrid Spain [email protected] E. E. Diplock Institute of Biological and Environmental Sciences Cruickshank Building University of Aberdeen Aberdeen, AB24 3UU UK

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Jayna L. Ditty Department of Biology University of St. Thomas St. Paul, MN 55105 USA [email protected] Jan Dolfing School of Civil Engineering and Geosciences Newcastle University Cassie Building 3.10 Newcastle Upon Tyne NE1 7RU UK [email protected] Patricia Domı´nguez-Cuevas Department of Environmental Protection Estaciόn Experimental del Zaidı´n, CSIC 18008 Granada Spain Rau´l Donoso Departamento de Gene´tica Molecular y Microbiologı´a Facultad de Ciencias Biolo´gicas NM-EMBA, CASEB, P. Universidad Cato´lica de Chile Alameda 340 Santiago Chile Vı´tor Martins dos Santos Systems and Synthetic Biology Group Helmholtz Centre for Infection Research (HZI) Inhoffenstrasse 7 38124 Braunschweig Germany [email protected] Gregory S. Douglas NewFields Environmental Forensic Practice LLC, Rockland MA 02370 USA [email protected]

Nicole Dubilier Symbiosis Group Max Planck Institute for Marine Microbiology Celsiusstrasse 28359 Bremen Germany [email protected]

Peter F. Dunfield Department of Biological Sciences University of Calgary 2500 University Dr. NW Calgary, Alberta Canada [email protected]

Estrella Duque Department of Environmental Protection Estaciόn Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain

Robert Duran Equipe Environnement et Microbiologie UMR CNRS IPREM 5254 Universite´ de Pau et des Pays de l’Adour (IPKEM-EEM), IBEAS BP1155 – UFR Sciences Avenue de l’Universite´ 64013 Pau Cedex France [email protected]

Thomas Ebensen Department of Vaccinology and Applied Microbiology Helmholtz Center for Infection Research Inhoffenstraße 7 38124 Braunschweig Germany [email protected]

List of Contributors

Virginia Edgcomb Department of Geology and Geophysics Woods Hole Oceanographic Institution Woods Hole MA 02543 USA [email protected] Rik IL Eggen Eawag, Swiss Federal Institute of Aquatic Science and Technology U¨berlandstrasse 133 8600 Du¨bendorf Switzerland [email protected] Thomas Egli Department of Environmental Microbiology, Eawag Swiss Federal Institute of Aquatic Science and Technology U¨berlandstrasse 133 8600 Du¨bendorf Switzerland [email protected] Lynda B. M. Ellis Department of Laboratory Medicine and Pathology Minneapolis MN 55455 USA [email protected] Matthias Engel Department of Biology University of Kaiserslautern 67663 Kaiserslautern Germany [email protected] Manuel Espinosa-Urgel Consejo Superior de Investigations Cientificas EEZ-CSIC Granada Spain

Anne Fahy Department of Biological Sciences University of Essex Central Campus Wivenhole Park Colchester C04 35Q UK [email protected]

Manuel Ferrer Institute of Catalysis CSIC, Centro de Investigaciones Biolo´gicas Marie Curie 2 28040 Madrid Spain [email protected]

James Gregory Ferry Department of Biochemistry and Molecular Biology The Pennsylvania State University 205 S. Freas Laboratory University Park PA 16802 USA [email protected]

Susanne Fetzner Institut fu¨r Molekulare Mikrobiologie und Biotechnologie Westfa¨lische Wilhelms-Universita¨t Mu¨nster Corrensstrasse 3 48149 Mu¨nster Germany [email protected]

Sandy Fillet Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas 18008 Granada Spain

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Janett Fischer Department of Bioremediation Helmholtz Centre for Environmental Research - UFZ Permoserstr. 15 04138 Leipzig Germany

Barbara Fuchs Department of Vaccinology and Applied Microbiology Helmholtz Center for Infection Research Inhoffenstraße 7 38124 Braunschweig Germany [email protected]

Antje Flieger Pathogenesis of Legionella Infection Research Group NG 5 Robert Koch-Institut Nordufer 20 Berlin Germany [email protected]

Roberta R. Fulthorpe Physical and Environmental Sciences University of Toronto at Scarborough 1265 Military Trail Toronto, ON Canada [email protected]

Julia Foght Department of Biological Sciences University of Alberta Biological Sciences Bldg., Room M 440 Edmonton, Alberta Canada [email protected]

Shoko Furuno Department of Environmental Microbiology UFZ Helmholtz Centre for Environmental Research Leipzig Germany

Michael Formolo Department of Biogeochemistry Max Planck Institute for Marine Microbiology Celsiusstrasse 1 28359 Bremen Germany [email protected]

Claude Gaillardin Microbiologie et Ge´ne´tique Mole´culaire AgroParisTech CNRS UMR2585 INRA UMR1238 Thiverval Grignon France [email protected]

Sofı´a Fraile Systems Biology Program Centro Nacional de Biotecnologı´a-CSIC Campus de Cantoblanco Madrid 28049 Spain

Tamara Galloway Ecotoxicology Research Group University of Exeter, Hatherly Laboratory Prince of Wales Road Exeter UK [email protected]

Benjamin Franken Institute of Molecular Enzyme Technology Heinrich-Heine University Duesseldorf Research Center Juelich 52426 Juelich Germany [email protected]

Alexander Galushko Max Planck Institute for Marine Microbiology Celsiusstraße 1 28359 Bremen Germany [email protected]

List of Contributors

Vanina Garcı´a Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain

Philippe Ge´rard INRA, CR Jouy, Domaine de Vilvert UR 910 Ecologie et Physiologie du Syste´me Digestif Jouy-en-Josas France [email protected]

J. L. Garcı´a Departamento de Microbiologı´a Molecular Centro de Investigaciones Biolo´gicas Consejo Superior de Investigaciones Cientı´ficas Ramiro de Maeztu Madrid Spain

Christoph Gertler School of Biological Sciences Bangor University Deiniol Road Bangor, Gwyned LL57 2UW UK [email protected]

Natalie J. Garton Department of Infection, Immunity and Inflammation University of Leicester Medical School Maurice Shock Building LE1 9HN Leicester UK [email protected]

Otto Geiger Centro de Ciencias Geno´micas Universidad Nacional Auto´noma de Me´xico Cuernavaca CP62210 Morelos Mexico [email protected]

Maria Genovese Instituto per l’Ambiente Marino Costiero (IAMC) - CNR Spianata San Raineri 86 98122 Messina Italy [email protected]

Birgit Geueke Eawag Du¨bendorf Switzerland

Azam Ghazi CSIC, Institute of Catalysis Department of Applied Biocatalysis Marie Curie 2 28049 Madrid Spain

Jack A. Gilbert Plymouth Marine Laboratory Prospect Place, The Hoe Plymouth, PL 3DH UK [email protected]

Patricia Godoy Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain

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Marisol Gonˇi-Urriza Equipe Environnement et Microbiologie, UMR CHRS Universite´ de Pau et des Pays de l’Adour IPREM 5254 IBEAS BP1155 Avenue de l’Universite´ 64013 Pau cedex France [email protected] Howard Goldfine Department of Microbiology University of Pennsylvania School of Medicine 425/6 Johnson Pavilion Philadelphia, PA 19104 USA [email protected] Olga V. Golyshina Environmental Microbiology Laboratory HZI-Helmholtz Centre for Infection Research Inhoffenstr. 7 38124 Braunschweig Germany [email protected] Peter N. Golyshin Environmental Microbiology Laboratory HZI-Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany and School of Biological Bangor University Gwynedd LL57 2UW UK [email protected] Virendra S. Gomase Department of Bioinformatics Padmashree Dr. D. Y. Patil University CBD Belapur Navi Mumbai India

Bernado Gonza´lez Departamento de Gene´tica Molecular y Microbiologı´a Facultad de Ciencias Biolo´gicas NM-EMBA, CASEB P. Universidad Cato´lica de Chile Alameda 340 Santiago Chile

Jose M. Gonza´lez Department of Microbiology University of La Laguna 38071 La Lagana Tenerife Spain [email protected]

Anna Gorbushina LBMPS, De´partement de Biologie Ve´ge´tale Universite´ de Gene´ve 30, quai Ernest-Ansermet 1211 Gene´ve 4 Suisse

Avdrey Gramain Department of Biological Sciences University of Essex Wivenhoe Park C04 3SQ Colchester UK

Hugo C. Gramajo Instituto de Biologı´a Molecular y Celular de Rosario (IBR-CONICET) Departamento de Microbiologı´a Facultad de Ciencias Bioquı´micas y Farmace´uticas Universidad Nacional de Rosario Suipacha, Rosario Argentina [email protected]

List of Contributors

Russel Grant Faculty of Health Life and Social Sciences Napier University Edinburgh EH10 5DT Scotland Neil D. Gray School of Civil Engineering and Geosciences and Institute for Research on the Environment and Sustainability Newcastle University NE1 7RU Newcastle Upon Tyne UK Stefan J. Green Department of Oceanography Florida State University Tallahassee, FL USA Charles W. Greer Environmental Microbiology National Research Council Biotechnology Research Institute (NRC-BRI) 6100 Royal Mount Avenue Montre´al, QC Canada [email protected] Frauke Greve Bioprocess Laboratory Institute of Process Engineering ETH Zurich Zurich Switzerland [email protected] Christian Griebler Institute of Groundwater Ecology Helmholtz Zentrum Mu¨nchen German Research Center for Environmental Health Ingolsta¨dter Landstrasse 1

85164 Neuherberg Germany Re´gis Grimaud Institut Pluridisciplinaire de Recherche en Environnement et Mate´riaux Equipe Environnement et Microbiologie UMR 5254 CNRS, IBEAS Universite´ de Pau et des Pays de l’Adour Pau France [email protected] Charlotte Grootaert Laboratory of Microbial Ecology and Technology (LabMET) Ghent University Coupure Links 653 Ghent 9000 Belgium Olaf Grundmann Max Planck Institute for Marine Microbiology 28359 Bremen Germany [email protected] Marı´a-Eugenia Guazzaroni Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas 18008 Granada Spain Salvatore Guglielmino Dipartimento di Scienze della Vita Universita´ di Messina Messina Italy Erich Gulbins Department of Molecular Biology University of Duisburg-Essen Hufelandstrasse 55 45122 Essen Germany [email protected]

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Robert P. Gunsalus Department of Microbiology and Molecular Genetics University of California 609 Charles Young Drive East, UCLA Los Angeles, CA 90095 USA [email protected]

Carlos A. Guzma´n Department of Vaccinology & Applied Microbiology Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany [email protected]

John D. Haddock Department of Microbiology Southern Illinois University 1125 Lincoln Drive Carbondale, IL 62901-6508 USA [email protected]

Jo Hagger Ecotoxicology Research Group University of Exeter Hatherly Laboratory Prince of Wales Road Exeter UK

Steven J. Hallam Department of Microbiology and Immunology University of British Columbia Life Sciences Centre 2552–2350 Health Sciences Mall Vancouver, BC V6T 1Z3 Canada [email protected]

Peter F. Hallin Center for Biological Sequence Analysis Technical University of Denmark 2800 Kgs. Lyngby Denmark John E. Hallsworth School of Biological Sciences MBC, Queen’s University Belfast Belfast, BT9 7BL Northern Ireland [email protected] Jens Harder Department of Microbiology Max Planck Institute for Marine Microbiology Bremen Germany [email protected] Hauke Harms Department of Environmental Microbiology UFZ Helmholtz Centre for Environmental Research Permoserstraße 15 04318 Leipzig Germany [email protected] Janet K. Hatt School of Civil and Environmental Engineering Georgia Institute of Technology 311 Ferst Drive Atlanta, GA USA Sascha Hausmann Institut fu¨r Molekulare Enzymtechnologie Heinrich-Heine-Universita¨t Du¨sseldorf Forschungszentrum Ju¨lich 52426 Ju¨lich Germany and Evocatal GmbH

List of Contributors

Merowingerplatz 1a 40225 Du¨sseldorf Germany [email protected] Rudolf Hausmann Institute of Engineering in Life Sciences Section of Technical Biology University of Karlsruhe Karlsruhe Germany [email protected] Terry C. Hazen Microbial Ecology and Environmental Engineering Lawrence Berkeley National Laboratory One Cyclotron Road Berkeley, CA USA [email protected] Ian M. Head School of Civil Engineering and Geosciences Newcastle University Newcastle upon Tyne NE1 7RU UK [email protected] Zhili He Institute for Environmental Genomics and Department of Botany and Microbiology University of Oklahoma Norman OK 73019 USA Brian P. Hedlund School of Life Sciences University of Nevada Las Vegas, NV 89154-4004 USA [email protected]

Johann Heider Laboratory for Microbiology University of Marburg Marburg Germany [email protected]

Mark L. Heinnickel Department of Plant and Microbial Biology University of California Berkeley, CA USA

Hermann J. Heipieper Department of Bioremediation Helmholtz Centre for Environmental Research - UFZ Permoserstrasse 15 04318 Leipzig Germany [email protected]

M. Antonia Henares-Molina Department of Environmental Protection Estacio´n Experimental del Zaidı´n, Consejo Superior de Investigaciones Cientı´ficas Granada Spain Phil Hendry CSIRO Wealth from Oceans Flagship North Ryde NSW 2113 Australia Christiane T. Hennessee Department of Molecular Biosciences and Bioengineering University of Hawaii East-West Road Honolulu, HI 96822 USA

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Patrick Ho¨hener Equipe Chimie de l’Environnement Continental Universite´ de Provence-CNRS 3 Place Victor Hugo - Case Marseille Cedex 3 France [email protected] Tori Ho¨ehler Exobiology Branch NASA Ames Research Center Moffett Field CA 94035 USA [email protected] Christof Holliger ENAC-ISTE-Laboratory for Environmental Biotechnology Ecole Polytechnique Fe´de´rale de Lausanne (EPFL) Lausanne Switzerland [email protected] Sueharu Horinouchi Department of Biotechnology Graduate School of Agriculture and Life Sciences University of Tokyo 1-1-1 Yayoi, Bunkyo-ka Tokyo 113-8657 Japan [email protected] L. Hoyles Food Microbial Sciences Unit Department of Pharmacy Food Biosciences and Chemistry University of Reading Whiteknights, Reading RG6 6AP UK [email protected]

Haiping Huang Petroleum Reservoir Group Department of Geology and Geophysics and Alberta Ingenuity Center for In situ Energy University of Calgary Calgary, AB T2N 1N4 Canada Wei E. Huang Kroto Research Institute The University of Sheffield Broad Lane Sheffield S3 7HQ UK Casey Hubert Max Planck Institute for Marine Microbiology Celsiusstraße 1 28359 Bremen Germany [email protected] Norbert Hu¨sers Institute of Waste Management and Contaminated Site Treatment Technische Universita¨t Dresden Pratzschwitzer Str. 15 61796 Pirna Germany [email protected] Jon Huisman Ministry of Transport and Water Public Works and Water Management RWS-Noordzee, Lange Kleiweg 34 2288 GK Rijswijk The Netherlands [email protected] Daniel Hunkeler Center for Hydroge´ology University of Neuchaˆtel Rue Emile Argand 11 Neuchaˆtel Switzerland [email protected]

List of Contributors

Karl-Erich Jaeger Institute of Molecular Enzyme Technology Heinrich-Heine University Duesseldorf Research Center Juelich 52426 Juelich Germany [email protected] Dieter Jahn Institut fu¨r Mikrobiologie Technical University Braunschweig Spielmannstr. 7 38106 Braunschweig Germany Martina Jahn Institut fu¨r Mikrobiologie Technical University Braunschweig Spielmannstr. 7 38106 Braunschweig Germany [email protected] Rakesh Jain Institute of Microbial Tachnology University of Kalyani Section 39A Chandigarh 160036 India [email protected] Graeme N. Jarvis New Zealand Trade and Enterprise Biotechnology & Agritechnology Sector 100 Willis Street/Level 15 The Majestic Centre Wellington New Zealand [email protected] Jose´ I. Jime´nez Department of Microbial Biotechnology Centro Nacional de Biotecnologı´a-Consejo Superior de Investigaciones Cientı´ficas 28049 Madrid Spain [email protected]

Anders R. Johnsen Department of Geochemistry Geological Survey of Denmark and Greenland Oster Voldgade 10 1350 Copenhagen Denmark [email protected] Dave R. Johnson Department of Fundamental Microbiology Baˆtiment Biophore, University of Lausanne Lausanne Switzerland [email protected] Andrew W. B. Johnston School of Biological Sciences University of East Anglia Norwich NR4 7TJ UK [email protected] Arjen Jonas BioDetection Systems BV, Kruislaan 1098SM Amsterdam The Netherlands D. Martin Jones School of Civil Engineering and Geosciences Newcastle University Newcastle upon Tyne NE1 7RU England UK Yves Jouanneau CEA, iRTSV, LCBM, and CNRS UMR 5249 17 rue des Martyrs Grenoble France [email protected]

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Alan Judd Alan Judd Partnership High Mickley Northumberland UK Howard Junca AG Biodegradation Helmholtz - Zentrum fu¨r Infektionsforschung Inhoffenstraße 7 38124 Braunschweig Germany Peter Ka¨mpfer Institut fu¨r Angewandte Mikrobiologie Justus-Liebig-Universita¨t Giessen Giessen Germany [email protected] Nicolas Kalogerakis Department of Environmental Engineering, Technical University of Crete Polytechneioupolis 73100 Chania Greece [email protected] Rainer Kalscheuer Howard Hughes Medical Institute and Albert Einstein College of Medicine Price Center 569 1301 Morris Pare Avenue Bronx NY USA [email protected] Yisheng Kang Department of Chemical Engineering University of California Berkeley, CA USA Forest M. Kaser Department of Plant and Microbial Biology, University of California Berkeley, CA USA

Chiaki Kato Exremobiosphere Research Centre Japan Agency for Marine-Earth Science and Technology, 2-15 Natsushima-cho Yokosuka 237-0061 Japan [email protected] Akitomo Kawasaki Faculty of Life Sciences University of Manchester Michael Smith Bldg Oxford Rd Manchester M13 9PT UK Jay D. Keasling Departments of Chemical Engineering and Bioengineering University of California at Berkeley 717 Potter Street Berkeley, CA USA [email protected] Christoph Keel Department of Fundamental Microbiology University of Lausanne Baˆtiment Biophore bureau 2310A Quartier UNIL-Sorge 1015 Lausanne Switzerland [email protected] Donovan P. Kelly Department of Biological Sciences University of Warwick Coventry UK [email protected] Michael A. Kertesz Faculty of Life Sciences University of Manchester Michael Smith Bldg, Oxford Rd Manchester M13 9PT UK [email protected]

List of Contributors

Thomas L. Kieft New Mexico Institute of Mining and Technology Socorro, New Mexico 87801 USA [email protected]

Hans-Peter E. Kohler Du¨bendorf Environmental Microbiology Eawag U¨berlandstr. 133 Du¨bendorf Switzerland [email protected]

Paul Kienhuis Ministry of Transport and Water Management Waterdienst 8200 AA Lelystad The Netherlands [email protected]

Harald Kolmar Clemens-Scho¨pf-Institute Department of Biochemistry Technical University Darmstadt Darmstadt Germany

Sang-Jin Kim Marine Biotechnology Research Centre Korea Ocean Research and Development Institute Sa 2 Dong 1270 Ansan 425-600, Seoul Korea [email protected]

Ralf Koppmann Department of Physics - Atmospheric Physics University of Wuppertal Gauss Strasse 20 42119 Wuppertal Germany [email protected]

Bronwyn M. Kirby Department of Biotechnology Institute for Microbial Biotechnology and Metagenomics University of the Western Cape Bellville, 7535 Cape Town South Africa

Irina A. Kosheleva Skryabin Institute of Biochemistry and Physiology of Microorganisms Russian Academy of Sciences Pushchino State University Pushchino 142290 Russia

Katrin Knittel Max Planck Institute for Marine Microbiology Celiusstr. 1 28359 Bremen Germany [email protected]

Oleg R. Kotsyurbenko Technical University Braunschweig Institute of Microbiology Spielmannstr. 7 38101 Braunschweig Germany [email protected]

Yosuke Koga University of Occupational and Environmental Health 9-14-20 Hinosato Munakata City 811-3425 Japan [email protected]

Tino Krell Department of Environmental Protection Estacio´n Experimental del Zaidı´n Profesor Albareda 1 18008 Granada Spain [email protected]

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Wolfgang E. Krumbein Geomicrobiology, ICBM Carl von Ossietzky University Oldenburg 2611 Oldenburg Germany [email protected] Matthias Ka¨stner Department for Environmental Biotechnology Helmholtz Center for Environmental Research - UFZ Leipzig Germany [email protected] Leonid A. Kulakov School of Biological Sciences and The QUESTOR Centre The Queen’s University of Belfast Belfast BT9 5AG, Northern Ireland UK Ohgew Kweon Division of Microbiology National Center for Toxicological Research Food and Drug Administration Jefferson, AR 72079 USA Kae Kyoung Kwon Marine Biotechnology Research Centre Korea Ocean Research and Development Institute Sa 2 Dong 1270 Ansan 425-600 Korea Jesu´s Lacal Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas 18008 Granada Spain

Jeremy H. Lakey Institute for Cell and Molecular Biosciences University of Newcastle upon Tyne Framlington Place Newcastle upon Tyne NE2 4HH UK [email protected] Carolyn M. C. Lam Systems and Synthetic Biology Group Helmholtz Centre for Infection Research Inhoffenstraße 7 38124 Braunschweig Germany [email protected] Mike J. Larkin School of Biological Sciences and The QUESTOR Centre The Queen’s University of Belfast 97 Lisburn Road Belfast, Northern Ireland UK [email protected] Stephen R. Larter Petroleum Reservoir Group Department of Geology and Geophysics and Alberta Ingenuity Center for In situ Energy University of Calgary Calgary, AB T2N 1N4 Canada J. R. Lawrence Environment Canada Saskatoon Saskatchewan Canada Sylvie Le Borgne Department of Process and Technology UAM-Cuajimalpa Artificios 40, Col. Miguel Hidalgo Mexico City Mexico [email protected]

List of Contributors

Mary Beth Leigh Department of Biology and Wildlife and Institute of Arctic Biology University of Alaska Fairbanks 902 Koyukuk Dr., Irving I Fairbanks, AK USA [email protected]

Frank Leitermann University of Karlsruhe Institute of Engineering in Life Sciences Section of Technical Biology Karlsruhe Germany

Ceri Lewis Ecotoxicology Research Group University of Exeter, Hatherly Laboratory Prince of Wales Road Exeter UK

Yuting Liang Institute for Environmental Genomics Department of Botany and Microbiology University of Oklahoma Norman, OK USA

Xiang Li Department of Molecular Biology University of Duisburg-Essen Hufelandstrasse 55 45122 Essen Germany Qing Xiao Li Molecular Biosciences and Bioengineering University of Hawaii 1955 East-West Road Honolulu, HI USA [email protected]

G. Li Institute for Environmental Genomics Department of Botany and Microbiology University of Oklahoma Norman, OK USA Mary E. Lidstrom Department of Chemical Engineering Microscale Life Sciences Center University of Washington Seattle, WA 98195–2180 USA [email protected] Susanne Liebner Institute for Biogeochemistry and Pollutant Dynamics (IBP) Federal Institute of Technology (ETH) Universita¨tstrasse 16 8092 Zu¨rich Switzerland [email protected] Keyu Liu CSIRO Wealth from Oceans Flagship 11 Julius Ave North Ryde NSW 2113 Australia Yuchen Liu Department of Microbiology University of Georgia 541 Biological Science Building Athens, GA 30602 USA [email protected] Frank E. Lo¨ffler School of Civil and Environmental Engineering Georgia Institute of Technology 311 Ferst Drive, ES4T Atlanta, GA USA [email protected]

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Angelina Lo Giudice Department of Animal Biology and Marine Ecology University of Messina Salita Sperone Messina Italy [email protected] N. Lo´pez-Corte´s Department of Applied Biocatalysis CSIC, Institute of Catalysis 28049, Madrid Spain Isabel M. Lo´pez-Lara Ecological Genomics Program Centre for Genomic Sciences Cuernavaca Morelos Me´xico Tillmann Lueders Institute of Groundwater Ecology Helmholtz Zentrum Mu¨nchen German Research Center for Environmental Health Ingolstaedter Landstr. 1 85764 Neuherberg Germany [email protected] Heinrich Lu¨nsdorf Environmental Microbiology Laboratory HZI-Helmholtz Centre for Infection Research 38124 Braunschweig Germany heinrich.luensdorf@helmholtz-hzide Boguslaw Lupa Department of Microbiology The University of Georgia 541 Biological Sciences Building Athens, GA 30602-2605 USA [email protected]

Alexandre J. Macedo Universidade Federal do Rio Grande do Sul Faculdade de Farma´cia and Centro de Biotecnologia Av. Ipiranga, 2752 Porto Alegre, RS Brazil Mark C. Malpass Bangor University School of the Environment and Natural Resources Deiniol Road Bangor, Gwyned LL572UW UK Roger Marchant School of Biomedical Sciences University of Ulster Coleraine, County Londonderry BT52 1SA Northern Ireland UK Silvia Marque´s Department of Environmental Protection Estacio´n Experimental del Zaidı´n, CISC Apartad 419 Granada Spain [email protected] Mariano A. Martı´nez Instituto de Biologı´a Molecular y Celular de Rosario (IBR-CONICET) Departamento de Microbiologı´a Facultad de Ciencias Bioquı´micas y Farmace´uticas Universidad Nacional de Rosario Suipacha, Rosario Argentina [email protected] Paula M. Martı´nez Department of Bioremediation Helmholtz Centre for Environmental

List of Contributors

Research-UFZ Permoserstr. 15 04318 Leipzig Germany J. L. Martı´nez Centro Nacional de Biotecnologı´a CSIC, Cantoblanco Madrid Spain Miguel Matilla Department of Environmental Protection EEZ-CSIC Granada Spain Monika Maurhofer Institute of Integrative Biology Plant Pathology Swiss Federal Institute of Technology (ETH) 8092 Zu¨rich Switzerland Erin M. McCammick School of Biological Sciences MBC, Queen’s University Belfast Belfast Northern Ireland Christopher W. J. McChalicher Department of Chemical Engineering and Materials Science University of Minnesota Minneapolis Minnesota USA Stephen McClean School of Biomedical Sciences University of Ulster Coleraine County Londonderry BT52 1SA Northern Ireland UK

Terry J. McGenity Department of Biological Sciences University of Essex Central Campus Wivonhole Park Colchester CO4 3SQ UK [email protected]

Michael J. McInerney Department of Botany and Microbiology University of Oklahoma 770 Van Vleet Oval Norman, OK USA [email protected]

Boyd McKew Department of Biological Sciences University of Essex Colchester CO4 35Q UK [email protected]

Rainer U. Meckenstock Institute of Groundwater Ecology Helmholtz Zentrum Mu¨nchen German Research Center for Environmental Health Ingolsta¨dter Landstraße 1 85764 Neuherberg Germany [email protected]

Farrakh Mehboob Wageningen University and Research Center Laboratory of Microbiology Dreijenplein 10 6703 HB Wageningen The Netherlands [email protected]

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Friedhelm Meinhardt Institut fu¨r Molekulare Mikrobiologie und Biotechnologie Westfa¨lische Wilhelms-Universita¨t Mu¨nster Corrensstr. 3 48149 Mu¨nster Germany Luigi Michaud Department of Animal Biology and Marine Ecology University of Messina Salita Sperone 31 Messina Italy [email protected] Akimasa Miyanaga Department of Biotechnology Graduate School of Agriculture and Life Sciences University of Tokyo 1-1-1 Yayoi Bunkyo-ku Tokyo 113-8657 Japan J. Michael Moldowan Department of Geological & Environmental Sciences Stanford University Stanford, CA USA [email protected] S. Monchy Brookhaven National Laboratory (BNL) Biology Department Upton, NY USA [email protected] Edward R. B. Moore Department of Clinical Bacteriology CCUG - Culture Collection University of Go¨teborg Sahlgrenska University Hospital Guldhedsgatan 10A

41346 Go¨teborg Sweden [email protected] Marcia Morales Department of Process and Technology UAM-Cuajimalpa Mexico City Mexico Susann Mu¨ller Helmholtz Centre for Environmental Research-UFZ Department of Environmental Microbiology, AG Flow Cytometry Permoserstraße 15 04318 Leipzig Germany Kai Muffler Institute of Bioprocess Engineering University of Kaiserslautern Kaiserslautern Germany Cormac D. Murphy UCD School of Biomolecular and Biomedical Science and Centre for Synthesis and Chemical Biology University College Dublin Belfield, Dublin 4 Ireland [email protected] J. Colin Murrell Department of Biological Sciences University of Warwick Coventry CV4 7AL UK [email protected] Florin Musat Max Planck Institute for Marine Microbiology Celsiusstraße 1 28359 Bremen Germany [email protected]

List of Contributors

Sine´ad M. Nı´ Chadhain Biotechnology Center for Agriculture and the Environment Cooks College Rutgers University New Brunswick, NJ USA M. Nachtkamp Institute for Chemistry and Biology of the Marine Environment (ICBM) University of Oldenburg 26111 Oldenburg Germany Gloria Navarro-Avile´s Department of Environmental Protection EEZ- CSIC Granada Spain Thomas R. Neu Helmholtz Centre for Environmental Research - UFZ Magdeburg Germany [email protected] Josh D. Neufeld Department of Biology University of Waterloo 200 University Avenue West Waterloo Ontario, N2L 391 Canada [email protected] Cecile Neuve´glise Laboratoire de Microbiologie et Ge´ne´tique Mole´culaire INRA, AgroParisTech Centre de Biotechnologie Agro-Industrielle INRA,UMR 1238, CNRS, UMR 2585 Thiverval-Grignon France

Jean-Marc Nicaud Laboratoire de Microbiologie et Ge´ne´tique Mole´culaire INRA, UMR 1238, CNRS, UMR 2585 AgroParisTech Centre de Biotechnologie AgroIndustrielle Thiverval-Grignon France [email protected] Nancy N. Nichols National Center for Agricultural Utilization Research Agricultural Research Service U.S. Department of Agriculture Peoria, IL USA [email protected] Kenneth W. Nickerson School of Biological Sciences University of Nebraska Lincoln, NE USA 68588-0666 [email protected] M. Nicolo` Dipartimento di Scienze della Vita Universita` di Messina Messina Italy Thomas D. Niederberger Biotechnology Research Institute National Research Council of Canada Montre´al QC Canada Jeppe L. Nielsen Department for Biotechnology Chemistry and Environmental Engineering Aalborg University Sohngaardsholmsvej 49 9000 Aalborg Denmark [email protected]

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Per H. Nielsen Department for Biotechnology Chemistry and Environmental Engineering Aalborg University Sohngaardsholmsvej 49 9000 Aalborg Denmark [email protected] Helge Niemann Max Planck Institute for Marine Microbiology Bremen, Germany and Institute for Environmental Geosciences University of Basel Bernioullistr. 30 Basel Switzerland [email protected]

Juan Nogales Department of Molecular Microbiology Centro de Investigaciones Biolo´gicasConsejo Superior de Investigaciones Cientı´ficas 28040 Madrid Spain Bernard Ollivier Laboratoire de Microbiologie, IRD, UMR 180 Universite´ de la Me´diterrane´e Faculte´ des Sciences de ESIL Cae 925 163 avenue de Luminy 13288 Marseille Cedex 9 France [email protected] Carrie O’Malley Department of Biology University of York Yo 10 5YW York UK

Maria Nikolopoulou Department of Environmental Engineering Technical University of Crete Polytechneioupolis 73100 Chania Greece [email protected]

Louise A. O’Sullivan Cardiff School of Biosciences Cardiff University Main Building Park Place Cardiff, Wales CF10 3TL UK

Julis R. Kemadjou Njiwa Eawag, Swiss Federal Institute of Aquatic Science and Technology U¨berlandstrasse 133 8600 Du¨bendorf Switzerland

Elizabeth Padilla-Crespo School of Civil Biology Georgia Institute of Technology 311 Ferst Drive Atlanta, GA 30332 USA

Balbina Nogales Department Biologia Universitat de les Illes Balears Crtra. Valldemossa Km 7.5 07122 Palma de Mallorca Spain [email protected]

S. Paı¨sse´ Equipe Environnement et Microbiologie UMR CNRS IPREM Universite´ de Pau et des Pays de l’Adour IBEAS Pau cedex France

List of Contributors

Norberto J. Palleroni Department of Biochemistry and Microbiology Rutgers University New Brunswick NJ, USA [email protected] Janmejay Pandey Institute of Microbial Technology University of Kalyani Chandigarh 160036 India [email protected] Sven Panke Bioprocess Laboratory Institute of Process Engineering ETH Zurich Universita¨tsstrasse 6 Zurich Switzerland [email protected] Rebecca E. Parales Department of Microbiology University of California 226 Briggs Hall 1 Shields Avenue Davis, CA 95616 USA [email protected] R. John Parkes School of Earth and Ocean Sciences Cardiff University Main Building, Park Place Cardiff, Wales CF10 3YE UK [email protected] Victor Parro Laboratory of Molecular Ecology, Centro de Astrobiologı´a (INTA-CSIC), Carretera de Ajalvir Km4 Torrejόnde Ardoz

Madrid Spain [email protected] Graeme Paton School of Biological Sciences University of Aberdeen Room 2.22, Cruickshank Building Aberdeen Scotland UK [email protected] Martina Pavlova Loschmidt Laboratories Institute of Experimental Biology and National Centre for Biomolecular Research Masaryk University Kamenice 5/A4 62500 Brno Czech Republic Maria Pe´chy-Tarr Department of Fundamental Microbiology University of Lausanne 1015 Lausanne Switzerland Ann Pearson Harvard University Department of Earth and Planetary Sciences 20 Oxford Street Cambridge, MA 02138 USA [email protected] Louis Peperzak Royal Netherlands Institute for Sea Research/NIOZ Landsdiep 4 1790 AB Den Burg Texel The Netherlands [email protected]

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Milva Pepi Department of Environmental Sciences University of Siena Via P.A. Mattioli 4 53100 Siena Italy Amedea Perfumo School of Biomedical Sciences University of Ulster Coleraine, County Londonderry BT52 1SA Northern Ireland UK Annelie Pernthaler Helmholtz Center for Environmental Research - UFZ Department of Environmental Microbiology Permoserstraße 15 04318 Leipzig Germany [email protected] Ken E. Peters U.S. Geological Survey 345 Middlerfield Road M5969 Menlo Park, CA USA [email protected] Jillian M. Petersen Symbiosis Group Max Planck Institute for Marine Microbiology Celsiusstrasse 1 28359 Bremen Germany [email protected] Erik Petrovskis Geosyntec Consultants 8120 Main Street Dexter, MI 48130 USA Dietmar H. Pieper Biodegradation Research Group Division of Microbiology Pathogenesis

HZI - Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany [email protected] Jo¨erg Pietruszka Institute of Bioorganic Chemistry Heinrich-Heine-University Duesseldorf Research Centre Juelich 52426 Juelich Germany [email protected] Cecilia Pini Department of Environmental Protection Estaciόn Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain Bradley Plantz School of Biological Sciences University of Nebraska Lincoln, NE 68588 - 0666 USA [email protected] Caroline M. Plugge Laboratory of Microbiology Wageningen University Dreijenplein 10 6703 HB Wageningen The Netherlands [email protected] Simon J. T. Pollard Centre for Resource Management and Efficiency Sustainable Systems Department School of Applied Sciences Cranfield University Cranfield, MK 43 OAL UK

List of Contributors

Danilo Pe´rez-Pantoja Departamento de Gene´tica Molecular y Microbiologı´a Facultad de Ciencias Biolo´gicas NM-EMBA, CASEB, P. Universidad Cato´lica de Chile Alameda 340 Santiago Chile Roger C. Prince ExxonMobil Biomedical Sciences, Inc. 1545 Route 22 East Annandale NJ 08801 USA [email protected] Giora Proskurowski Woods Hole Oceanographic Institution Department of Marine Chemistry and Geochemistry 266 Woods Hole Rd Woods Hole, MA 02543 USA [email protected] Jacek Puchałka Systems and Synthetic Biology Group Helmholtz Centre for Infection Research Inhoffenstraße 7 38124 Braunschweig Germany Alfred Pu¨hler Department of Genetics Bielefeld University 33594 Bielefeld Germany Jian G. Qin School of Biological Sciences Flinders University GPO BOX 2100 Adelaide 5001 Australia [email protected]

Ralf Rabus Institute for Chemistry und Biology of the Marine Environment (ICBM) Carl von Ossietzky University 26110 Oldenburg Germany [email protected] Juan L. Ramos Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Profesor Albaseda, 1 18008 Granada Spain [email protected] Maria-Isabel Ramos-Gonza´lez Department of Environmental Protection EEZ-CSIC Granada Spain Daniel S. Read Centre for Ecology and Hydrology Oxford Mansfield Road Oxford, OX1 3SR UK James W. Readman Plymouth Marine Laboratory Prospect Place, The Hoe Plymouth Devon UK [email protected] Walter Reineke Chemical Microbiology Bergische Universita¨t Wuppertal Gaußstr. 20 42097 Wuppertal Germany

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Friedrich Reinhard Department of Fundamental Microbiology University of Lausanne Baˆtiment Biophore Quartier UNIL-Sorg 1015 Lausanne Switzerland Oleg N. Reva Biochemistry Department Bioinformatics and Computational Biology Unit University of Pretoria Hillerest Pretoria South Africa [email protected] Hans H. Richnow Department for Isotope Biogeochemistry Helmholtz Center for Environmental Research - UFZ Permoserstrasse 15 04318 Leipzig Germany [email protected] Kirsti M. Ritalahti School of Civil and Environmental Engineering Georgia Institute of Technology 311 Ferst Drive Atlanta, GA 30332 USA Sagrario Arias Rivas Environmental Microbiology Laboratory Helmholtz Centre for Infection Research Inhoffenstrasse 7 38124 Braunschweig Germany [email protected] Frank T. Robb Center of Marine Biotechnology Biotechnology Institute

University of Maryland Baltimore, MD 21202 USA Laure`ne Rochat Department of Fundamental Microbiology University of Lausanne 1015 Lausanne Switzerland

Sara Rodrı´guez-Conde Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain Jose´ J. Rodrı´guez-Herva Department of Environmental Protection EEZ- CSIC Granada Spain Wilfred F. M. Ro¨ling Department of Molecular Cell Physiology Faculty of Earth and Life Sciences VU University Amsterdam De Boelelaan 1085 1081 HV Amsterdam and The Netherlands and NGI Ecogenomics Cluster Amsterdam The Netherlands [email protected] Michel Rohmer Institut Le Bel Universite´ Louis Pasteur–CNRS 4 rue Blaise Pascal 67070 Strasbourg France [email protected]

List of Contributors

Fernando Rojo Centro Nacional de Biotecnologı´a CSIC C/Darwin 3, Campus UAM, Cantoblanco 28049 Madrid Spain [email protected] Eliora Ron Department of Molecular Microbiology and Biotechnology Tel-Aviv University Ramat Tel-Aviv Israel [email protected] Jean-Franc¸ois Rontani Faculte des Sciences de Luminy Centre de’Oce´anologie de Marseille et de Bioge´ochimie (UMR 65359) CASE 901 Marseille France [email protected] Antonio Rosato Magnetic Resonance Center (CERM) -University of Florence Sesto Fiorentino Italy [email protected] Frank Rosenau Institute for Molecular Enzyme Technology Heinrich-Heine-University Duesseldorf Stetternicher Forst Ju¨lich Germany [email protected] Eugene Rosenberg Department of Molecular Microbiology and Biotechnology Tel-Aviv University Ramat Tel Aviv

Israel [email protected] Michael Rother Institut fu¨r Molekulare Biowissenschaften Molekulare Mikrobiologie & Bioenergetik Johann Wolfgang Goethe-Universita¨t Campus Rieddberg Max-von-Laue-Strasse 9 60438 Frankfurt am Main Germany [email protected] Arlene K. Rowan School of Civil Engineering and Geosciences Newcastle University NE1 7RU Newcastle upon Tyne UK Julia Sabirova Laboratory of Industrial Microbiology and Biocatalysis Department of Biochemical and Microbial Technology Faculty of Bioscience Engineering Ghent University Coupure L. 653 9000 Ghent Belgium [email protected] Flavia Talarico Saia Wageningen University and Research Center Laboratory of Microbiology Dreijenplein 10 6703 HB Wageningen The Netherlands Andrea Salis Dipartimento di Scienze Chimiche Universita` di Cagliari - CSGI Cittadella Universitaria Monserrato SS 554 Divio Sastu Monserrato (CA) Italy

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Paı¨sse Sandrine Equipe Environment et Microbiologie UMR CNRS IPREM 5254 Universite´ de Pau et des Pays de l’Adour, IBEAS BP 1155 Avenue de l’Universite´ 64013 Pau Cedex France Henrik Sass School of Earth and Ocean Sciences Cardiff University Main Building Park Place Cardiff, Wales CF 10 3YE UK Judith M. Schicks Helmholtz Centre Potsdam - GFZ German Research Centre for Geosciences Section 4.2 Telegrafenberg Haus B127 14473 Potsdam Germany [email protected] Bernhard Schink Department of Biology University of Konstanz 78457 Konstanz Germany [email protected] Andreas Schmid Institute for Analytical Sciences (ISAS) Bunsen-Kirchhoff-Strasse 11 44139 Dortmund Germany [email protected] Susanne Schneiker-Bekel Center for Biotechnology (CeBiTec) Bielefeld University 33594 Bielefeld Germany Max Schobert Institute of Microbiology Technische Universita¨t Braunschweig

Spielmannstr. 7 38106 Braunschweig Germany [email protected] Gosse Schraa Wageningen University and Research Center Laboratory of Microbiology Dreijenplein, Wageningen The Netherlands [email protected] Gustavo E. Schujman Instituto de Biologı´a Molecular y Celular de Rosario (IBR-CONICET) Departamento de Microbiologı´a Facultad de Ciencias Bioquı´micas y Farmace´uticas Universidad Nacional de Rosario Suipacha 531 2000 Rosario Argentina [email protected] Kai Schulze Department of Vaccinology and Applied Microbiology Helmholtz Center for Infection Research Inhoffenstraße 7 38124 Braunschweig Germany [email protected] Lorenz Schwark Institute for Geosciences Christian-Albrechts-University Ludewig-Meyn-Str. 10 24118 Kiel Germany [email protected] Jan Schwarzbauer Institute of Geology and Geochemistry of Petroleum and Coal RWTH Aachen University Lochnerstraße 4-20 52056 Aachen Germany [email protected]

List of Contributors

Michael Seeger Laboratorio de Microbiologı´a Molecular y Biotecnologı´a Ambiental Millennium Nucleus of Microbial Ecology and Environmental Microbiology and Biotechnology Departamento de Quı´mica, Universidad Te´cnica Federico Santa Marı´a Avenida Espan˜a 1680 Valparaı´so Chile [email protected]

Ana Segura Department of Environmental Protection Estaciόn Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain Drazˇenka Selesi Institute of Groundwater Ecology Helmholtz Zentrum Munchen German Research Center for Environmental Health Ingolsta¨dter Landstraße 85764 Neuherberg Germany Zongze Shao Key Laboratory of Marine Biogenetic Resources The Third Institute of Oceanography State Oceanic Administration Xiamen 361005, Fujian China [email protected] Angela Sherry Institute for Research on the Environment and Sustainability Newcastle University NE1 7RU Newcastle Upon Tyne UK

Victoria Shingler Department of Molecular Biology Umea˚ University 901 87 Umea˚ Sweden [email protected] Jessica R. Sieber Department of Botany and Microbiology University of Oklahoma 770 Van Vleet oval Norman, OK 73019 USA [email protected] Hortencia Silva-Jime´nez Department of Environmental Protection Estacio´n Experimental del Zaidίn Consejo Superior de Investigaciones Cientίficas 18008 Granada Spain Hauke Smidt Laboratory of Microbiology Wageningen University Dreijenplein 10 6703 HB Wageningen The Netherlands Kilian E. C. Smith Department of Environmental Chemistry and Microbiology National Environmental Research Institute (NERI) Aarhus University Frederiksborg kej 399 4000 Roskilde Denmark [email protected] Cindy J. Smith Department of Animal and Plant Sciences The University of Sheffield Western Bank Sheffield TN UK [email protected]

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Thomas J. Smith Biomedical Research Centre Sheffield Hallam University Howard Street Sheffield SI 1WB UK Thomas J. P. Smyth School of Biomedical Sciences University of Ulster Coleraine County Londonderry BT52 ISA Northern Ireland UK Christian Sohlenkamp Centro de Ciencias Geno´micas Universidad Nacional Auto´noma de Mexico Morelos CP62210 Me´xico Vincenzo Solinas Dipartimento di Scienze Chimiche Universita` di Cagliari - CSGI Cittadella Universitaria Monserrato 09042 Monserrato Italy Edwin Sonneveld BioDetection Systems BV Kruislaan Badhuisweg 3 1031 Amsterdam The Netherlands [email protected]

Minneapolis, St. Paul USA [email protected] James T. Staley Department of Microbiology University of Washington Seattle, WA 98195 [email protected] Alfons J. M. Stams Laboratory of Microbiology Wageningen University Dreijenplein 10 6703 HB Wageningen The Netherlands [email protected] Thorsten Stoeck Department of Biology Emmy-Noether Research Group “Marine Microcukaryotic Diversity” University of Kaiserslautern Erwin-Schro¨dinger Str. 4 67663 Kaiserslautern Germany [email protected] Marc Strous Max Planck Institute for Marine Microbiology Celsiusstrasse 25839 Bremen Germany [email protected]

Diana Z. Sousa The Netherlands Institute for Biotechnology and Bioengineering Centre of Biological Engineering University of Minho Braga Portugal

Sujatha Subramoni Bacteriology Group International Centre for Genetic Engineering & Biotechnology Treviso Italy [email protected]

Friedrich Srienc Department of Chemical Engineering and Materials Science University of Minnesota

Erwin Suess Leibniz-Institute for Marine Sciences (IFM-GEOMAR) Kiel and German Marine Research Consortium

List of Contributors

(KUM) 24148 Berlin Germany [email protected] Zulma R. Sua´rez-Moreno Bacteriology Group International Centre for Genetic Engineering & Biotechnology Padriciano 99 34012 Treviso Italy [email protected] Marc J.-F. Suter Eawag Swiss Federal Institute of Aquatic Science and Technology Ueberlandstrasse 133 8600 Dubendorf Switzerland John B. Sutherland Division of Microbiology National Center for Toxicological Research Food and Drug Administration Jefferson, AR 72079 USA [email protected] Christoph Syldatk Institute of Engineering in Life Sciences Institute of Bio- and Food Technology University of Karlsruhe Kaiserstr. 12 76131 Karlsruhe Germany [email protected] Safiyh Taghavi Brookhaven National Laboratory (BNL) Biology Department Upton, NY 11973-5000 USA [email protected] Marcus Taupp Department of Microbiology and Immunology

University of British Columbia Life Sciences Centre 2552-2350 Health Sciences Mall Vancouver, BC V6T 1Z3 Canada Lee Taylor Institute of Arctic Biology University of Alaska Fairbanks Fairbanks, AK 99775-7000 USA [email protected] Robin Tecon Department of Fundamental Microbiology University of Lausanne Baˆtiment Biophore Quartier UNIL-Sorge 1015 Lausanne Switzerland Eva Teira Departamento Ecoloxı´a e Bioloxı´a Animal Universidade de Vigo Campus Lagoas-Marcosende 36310 Vigo Spain [email protected] Wilson Tera´n Department of Biology Universidad Javeriana Bogota´ Colombia Andreas Teske Department of Marine Sciences University of North Carolina at Chapel Hill 351 Chapman Hall, CB# 3300 Chapel Hill USA [email protected] David Theaker Houghton plc Trafford Park Manchester M17 1AF UK [email protected]

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France Thevenieau Laboratoire de Microbiologie et Ge´ne´tique Mole´culaire INRA, UMR 1285, CNRS, UMR 2585 AgroParisTech Centre de Biotechnologie AgroIndustrielle BP 01, 78850 Thiverval-Grignon France and Oxyrane UK Limited, Greenheys House Manchester Science Park 10 Pencroft Way Manchester M15 655 UK Ian P. Thompson Department of Engineering Science Institute of Advanced Technologies University of Oxford Begbroke Science Park, Sandy Lane Yarnton OX51PF UK [email protected] Steven F. Thornton Groundwater Protection and Restoration Group Kroto Research Institute The University of Sheffield Sheffield S3 7HQ UK [email protected] Meghan Tierney Biotechnology Center for Agriculture and the Environment Rutgers University New Brunswick, NJ 08901 USA Kenneth N. Timmis Environmental Microbiology Laboratory Helmholtz Centre for Infection Research Inhoffenstrasse 7

38124 Braunschweig Germany David J. Timson School of Biological Sciences MBC, Queen’s University Belfast Belfast Northern Ireland Nils Tippkotter Institute of Bioprocess Engineering University of Kaiserslautern Kaiserslautern Germany Eva M. Top Department of Biological Sciences Initiative for Bioinformatics and Evolutionary Studies (IBEST) University of Idaho 258 Life Sciences Building South Moscow, ID USA [email protected] Jesu´s de la Torre Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas Granada Spain Kathleen Trautwein Max Planck Institute for Marine Microbiology 28359 Bremen Germany [email protected] Y. A. Trotsenko G.K.Skryabin Institute of Biochemistry and Physiology of Microorganisms Russian Academy of Sciences Pushchino Moscow Region Russia

List of Contributors

Burkhard Tu¨mmler Klinische Forschergruppe OE 6711 Medizinische Hochschule Hannover Hannover Germany [email protected] I. Marla Tuffin Center of Marine Biotechnology Biotechnology Institute University of Maryland Baltimore, MD USA [email protected] Roland Ulber Institute of Bioprocess Engineering University of Kaiserslautern Kaiserslautern Germany [email protected] David W. Ussery Center for Biological Sequence Analysis Technical University of Denmark 2800 Lyngby Denmark [email protected] William Ussler III Monterey Bay Aquarium Research Institute 7700 Sandholdt Road Moss Landing, CA 95039 USA Marc van Bemmel Bioclear BV Rozenburgloan 13 Groningen The Netherlands [email protected] Tom van de Wiele Laboratory of Microbial Ecology and Technology (LabMET) Ghent University Coupure L653

9000 Ghent Belgium [email protected] Bart van der Burg BioDetection Systems BV Kruislaan 1098 SM Amsterdam The Netherlands [email protected] Chris J. van der Gast NERC Centre for Ecology & Hydrology NERC Mansfield Road Oxford UK [email protected] Daniel van der Lelie Brookhaven National Laboratory (BNL) Biology Department Upton, NY USA [email protected] Jan Roelof van der Meer Department of Fundamental Microbiology University of Lausanne Baˆtiment Biophore, Quartier UNIL-Sorge 1015 Lausanne Switzerland [email protected] Pieter van Dillewijn Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas 18008 Granada Spain Jean van Heijenoort Institute of Biochemistry and Molecular and Cellular Biophysics University Paris-Sud Orsay 91405 France [email protected]

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Joy D. Van Nostrand Institue for Environmental Genomics and Department of Botany and Microbiology University of Oklahoma 101 David L. Boren Blvd. Norman, OK USA [email protected] J. Vangronsveld Hasselt University Department of Environmental Biology CMK, Universitaire 3590 Diepenbeek Belgium [email protected] Pierre-Joseph Vaysse Institut Pluridisciplinaire de Recherche en Environnement et Mate´riaux Equipe Environnement et Microbiologie UMR 5254 CNRS, IBEAS Universite´ de Pau et des Pays de l’Adour BP1155 64013 Pau France R. Vazquez-Duhalt Department of Cellular Engineering and Biocatalysis Instituto de Biotecnologia UNAM, Cuernavaca Mexico Caryn J. Vengadajellum Department of Chemical Engineering Biocatalysis and Technical Biology Unit University of Cape Town Rondebosch 7700 Cape Town South Africa Vittorio Venturi Bacteriology Group International Centre for Genetic Engineering & Biotechnology Padriciano 99

34012 Treviso Italy [email protected] Willy Verstraete Laboratory of Microbial Ecology and Technology (LabMET) Ghent University Coupure L653 9000 Ghent Belgium [email protected] Jose´ Maria´ Vieites CSIC, Institute of Catalysis 28049 Madrid Spain Andrea Vieth GeoForschungsZentrum Potsdam Organic Geochemistry Telegrafenberg, B227 14473 Potsdam Germany [email protected] Maria Vila-Costa Department of Marine Sciences University of Georgia Athens, GA 30602 USA [email protected] Christelle Vogne Department of Fundamental Microbiology University of Lausanne Lausanne Switzerland Herbert Volk CSIRO Wealth from Oceans Flagship 11 Julius Ave 2113 2113 North Ryde NSW Australia [email protected]

List of Contributors

Lawrence P. Wackett Department of Biochemistry, Molecular Biology, and Biophysics and Biotechnology Institute University of Minnesota 1479 Gortner Avenue St. Paul, MN 55108 USA [email protected] Marlea Wagelmans Bioclear BV Rozenburglaan 13 Groningen The Netherlands [email protected] Dirk Wagner Geomicrobiology and Carbon Dynamics in Periglacial Environments Alfred Wegener Institute for Polar and Marine Research Research Unit Potsdam Telegrafenberg A45 (Building A45-108) 14473 Potsdam Germany [email protected] John Wallace Gut Health Division Rowett Research Institute Bucksburn Aberdeen AB21 9SB UK [email protected] Vanisa Walter Institute of Engineering in Life Sciences Section of Technical Biology University of Karlsruhe Karlsruhe Germany Cliff Walters Corporate Strategic Research ExxonMobil Corporate Strategic Research 1545 Route 22 East

Annandale, NJ USA [email protected] Frederick J. Warren King’s College London Franklin Wilkins Building 150 Stamford Street London SEI GNH UK Trudy M. Wassenaar Center for Biological Sequence Analysis Technical University of Denmark 2800 Lyngby Denmark and Molecular Microbiology and Genomics Consultants 55576 Zotzenheim Germany [email protected] Kazuya Watanabe Research Center for Advanced Science and Technology University of Tokyo Tokyo Japan [email protected] Andrew J. Watkins School of Earth and Ocean Sciences Cardiff University Main Building Park Place Cardiff, Wales CF10 3YE UK Gordon Webster School of Earth and Ocean Sciences Cardiff University Main Building Park Place Cardiff, Wales CF 10 3YE UK

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Sander Weelink Wageningen University and Research Center Laboratory of Microbiology Dreijenplein 10 6703 HB Wageningen The Netherlands

Andrew S. Whiteley Biodiversity and Ecosystem Function Group, Molecular Microbiology Ecology Section CEH, Mansfield Road Oxford UK [email protected]

Andrew J. Weightman Cardiff School of Biosciences Cardiff University Main Building Park Place Cardiff, Wales CF 10 3TL UK

Lyle G. Whyte Department of Natural Resource Sciences McGill University Ste-Anne-de-Bellerne QC Canada

Frank Wenzho¨fer HGF MPG Joint Research Group on Deep Sea Ecology and Technology Alfred Wegener Institute for Polar and Marine Research Bremerhaven Germany

Lukas Y. Wick Department of Environmental Microbiology UFZ Helmholtz Centre for Environmental Research Permoserstraße 15 04318 Leipzig Germany [email protected]

Peter Werner Institute of Waste Management and Contaminated Site Treatment Technische Universita¨t Dresden Pratzschwitzer Str. 15 Dresden Germany [email protected] Nele Weyens Department of Environmental Biology Hasselt University 3590 Diepenbeek Belgium [email protected] C. Whitby Department of Biological Sciences University of Essex Colchester UK [email protected]

Fritz Widdel Max Planck Institute for Marine Microbiology Celsiusstraße 1 28359 Bremen Germany [email protected] Susanne Wilhelm Institute for Molecular Enzyme Technology Heinrich-Heine-University Du¨sseldorf Ju¨lich Germany Heinz Wilkes Helmholtz Centre Potsdam GFZ German Research Centre for Geosciences

List of Contributors

Organic Geochemistry Haus B22B Telegrafenberg 14473 Potsdam Germany [email protected] Ryan D. Wilson Groundwater Protection and Restoration Group Department of Civil and Structural Engineering University of Sheffield North Campus, Broad Lane Sheffield UK [email protected] Y. Witte Affiliations Royal Netherlands Institute for Sea Research (NIOZ) 1790 AB Den Burg Texel The Netherlands Rolf-M. Wittich Department of Environmental Protection Estacio´n Experimental del Zaidı´n Consejo Superior de Investigaciones Cientı´ficas 18008 Granada Spain Ann P. Wood Department of Microbiology King’s College London Dental Institute Floor 17 Guy’s Tower Guy’s Campus London SE1 9RT UK [email protected]

Liyou Wu Institute for Environmental Genomics and Department of Botany and Microbiology

University of Oklahoma Norman, OK 73019 USA Michail M. Yakimov Department of Earth and Environment Institute for Coastal Marine Environment CNR (National Research Council) Spinata S. Raineri 86 98122 Messina Italy [email protected] Lily Y. Young Biotechnology Center for Agriculture and the Environment Rutgers University Foran Hall, 59 Dudky Rd. New Brunswick, NJ 08901 USA [email protected] Yang Zhang Department of Molecular Biology University of Duisburg-Essen Hufelandstrasse 55 45122 Essen Germany Kun Zhang Jacobs School of Engineering University of California San Diego La Jolla CA 92093-0403 USA Jizhong Zhou Institute for Environmental Genomics and Department of Botany and Microbiology University of Oklahoma 101 David L. Boren Blvd. Norman, OK 73019 USA [email protected]

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Stephen H. Zinder Department of Microbiology Cornell University 272 Wing Hall Ithaca, NY 14850 USA [email protected] Smita Zinjarde Institute of Bioinformatics and Biotechnology University of Pune

Pune 4110007 India

Gerben J. Zylstra Biotechnology Center for Agriculture and the Environment Rutgers University Cooks College New Brunswick, NJ USA [email protected]

Part 1

Diversity and Physico-Chemical Characteristics

1 Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence H. Wilkes1 . J. Schwarzbauer2 1 Helmholtz Centre Potsdam, GFZ German Research Centre for Geosciences, Organic Geochemistry, Potsdam, Germany [email protected] 2 Institute of Geology and Geochemistry of Petroleum and Coal, RWTH Aachen University, Aachen, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5

2

Covalent Bonding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6

3 3.1 3.2 3.3 3.4

Saturated Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 n-Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Branched Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Cycloalkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Reactions of Saturated Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

4 Unsaturated Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 4.1 Alkenes and Alkines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 4.2 Reactions of Unsaturated Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16 5 5.1 5.2 5.3 5.4

Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 Aromaticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 Benzene Derivatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 Polycyclic Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 Reactions of Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21

6 Functionalized Organic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 6.1 Halogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 6.2 Oxygen and Sulfur . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_1, # Springer-Verlag Berlin Heidelberg, 2010

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6.3 Nitrogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29 6.4 Specific Reactions of Functionalized Organic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . 32 7

Bio- and Geomacromolecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36

8

Physical Properties of Hydrocarbons and Other Organic Compounds . . . . . . . . . . . . . 41

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

1

Abstract: Hydrocarbons are among the most abundant organic compound classes in the biogeosphere. They are formed directly by living organisms as biosynthetic products or through geological transformation of biomass in sedimentary systems. This article provides an introduction to the structural variability of hydrocarbons and their occurrence in natural environments. Besides saturated, unsaturated and aromatic hydrocarbons also selected types of functionalized organic compounds which play key roles in biogeochemical processes are discussed. For each compound type reactivity and important reaction types with a special focus on mechanisms relevant in biochemical transformations are presented. Bio- and geomacromolecules and their role in the formation of fossil fuels are briefly introduced. Important physico-chemical parameters are discussed in relation to the structural characteristics of the presented compound classes.

1

Introduction

Hydrocarbons occur in a great structural diversity as biosynthetic products of living organisms in the biosphere or as abiotic transformation products of biogenic organic matter in the geosphere. They are the main constituents of petroleum and thus are extremely abundant in geological systems. Increasing exploitation of hydrocarbon-based energy resources was one of the driving forces of the industrial revolution with dramatic impact on the evolution of human culture (e.g., Hall et al., 2003). The presence of hydrocarbons already on the Early Earth has promoted the evolution of metabolic pathways which allow microorganisms to exploit them as energy sources as well. The interactions of microorganisms and hydrocarbons are manifold. Hydrocarbons by definition contain exclusively the elements carbon and hydrogen. This chapter attempts to provide a compact introduction to fundamental aspects of hydrocarbon structure, properties, and occurrence which are most relevant for environmental and microbiological processes. It is unnecessary to say that it cannot substitute a textbook on organic chemistry. Hydrocarbons are divided into three main compound classes, namely (1) saturated, (2) unsaturated, and (3) aromatic hydrocarbons which are discussed separately in subsequent sections of this article. In addition to hydrocarbons sensu stricto this chapter also provides an introduction to important types of functionalized organic compounds containing the halogens fluorine, chlorine, bromine, or iodine or the hetero elements nitrogen, sulfur, and oxygen (NSO compounds). Halogenated organic compounds occurring in the environment are mainly released by anthropogenic activity although numerous natural products containing halogen atoms are known. NSO compounds may occur in natural environments along with complex assemblages of hydrocarbons, e.g., crude oils always contain nonhydrocarbons in highly variable but often significant amounts. Moreover, functionalized organic compounds, particularly oxygen compounds, represent important biological transformation products of hydrocarbons and play a crucial role as intermediates/products of biodegradation pathways. The discussion will consider both natural products as well as xenobiotics, i.e., compounds found in organisms which are not produced by these organisms or expected to occur in them. We will not give any introduction to the nomenclature of organic compounds. The common rules of organic nomenclature as defined by the International Union of Pure and Applied Chemistry (IUPAC) are accessible via suitable resources (Nomenclature of Organic Chemistry, Sections A, B, C, D, E, F, and H, Pergamon Press, Oxford, 1979. Copyright 1979 IUPAC; A Guide to IUPAC Nomenclature of Organic Compounds (Recommendations 1993),

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1993, Blackwell Scientific publications, Copyright 1993 IUPAC; http://www.acdlabs.com/ iupac/nomenclature/). Furthermore, software packages are available which generate the correct names of organic compounds from drawn structures (http://www.acdlabs.com/products/ name_lab/).

2

Covalent Bonding

A characteristic feature of hydrocarbons is that they contain covalent bonds exclusively. In covalent bonds, pairs of electrons are shared between atoms. Covalent bonds are typically formed between elements which do not differ too strongly in electronegativity (see below). The quantum mechanical valence bond model describes the nature of covalent bonds based on the assumption that atom orbitals overlap to form molecule orbitals which contain the shared electrons. Orbitals are regions around a single atom or in a molecule in which electrons may be found. In atoms of elements of the 2nd period of the periodic table of the chemical elements, such as carbon, nitrogen, oxygen, and fluorine, only atom orbitals of the 1st and 2nd shell, i.e., the 1s, 2s, 2px, 2py, and 2pz orbitals may be occupied by electrons. 1s, 2s, and 2p orbitals correspond to particular increasing energy levels of the electrons. Central to the understanding of covalent bonding is the concept of orbital hybridization, which assumes that mixing of energetically different atom orbitals forms energetically equivalent molecular orbitals. The three modes of orbital hybridization that may occur in carbon form four sp3, three sp2 or two sp orbitals by mixing of three, two or one 2p orbital(s) with the 2s orbital, respectively. Covalent bonds formed by overlapping of a sp3, sp2 or sp hybrid orbital with the 1s orbital of a hydrogen atom or a hybrid orbital of another carbon atom (or an atom of another element) are termed s-bonds. Carbon atoms with a sp3 hybridization form four single bonds along the connecting lines to the bonding partners. Methane (1), the simplest organic molecule, is build up by four equivalent C–H s-bonds. The structure of methane is that of a regular tetrahedron in which all bond lengths (110 pm) and bond angles (109.5
) are identical (> Fig. 1). The length of a C–C s-bond between two sp3 hybridized carbon atoms as in ethane (2) is 154 pm (> Fig. 1). The A-C-B bond angles in tetrahedral carbon atoms may deviate from 109.5
depending on the electronic and sterical properties of the bonding partners. Varying lengths (150–120 pm) are found for s-bonds between differently hybridized carbon atoms.

. Figure 1 Structures of simple hydrocarbons; (1), methane CH4; (2), ethane C2H6; (3), ethene C2H4; (4), ethine C2H2.

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Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

. Table 1 Elements most relevant to organic chemistry and biochemistry ordered by atomic number Element

H

C

N

O

F

P

S

Cl

Br

I

Atomic number

1

6

7

8

9

15

16

17

35

53

Electronegativity

2.20

2.55

3.04

3.44

3.98

2.19

2.58

3.16

2.96

2.66

Three or two s-bonds are formed by sp2 and sp hybridized carbon atoms, respectively. The remaining nonhybridized 2p orbital(s) overlap(s) with (a) nonhybridized 2p orbital(s) of other atoms to form p-bonds in addition to the s-bond. In p-bonds the molecular orbital is located above and below the connecting line between the atoms. Bonding between two sp2 or two sp hybridized carbon atoms thus leads to C–C double or triple bonds, respectively. Simple examples are ethene (3) (H2C=CH2) with a C–C double bond length of 134 pm and ethine (4) (HCCH) with a C–C triple bond length of 120 pm (> Fig. 1). The bonding angles at sp2 and sp hybridized carbon atoms are 120
and 180
, respectively. Carbon atoms may also form covalent bonds to atoms of elements other than carbon and hydrogen. Most relevant in naturally occurring organic compounds are halogen, nitrogen, sulfur, and oxygen (> Table 1). As the halogens are monovalent elements, between carbon and halogen atoms only single bonds are possible. In contrast, oxygen and sulfur are divalent elements and nitrogen is a trivalent element. Therefore, C-O and C-S single and double bonds as well as C–N single, double, and triple bonds are possible. The chemical reactivity of organic compounds depends directly on the properties of the individual bonds within their molecules. The strength of a bond is measured by the bond dissociation energy which is directly related to the bond distance. In chemical reactions, bond cleavage may occur by homolytic or heterolytic mechanisms. An example of a homolytic mechanism would be the cleavage of an alkane into an alkyl radical and a hydrogen radical, i.e., both cleavage products retain one of the shared electrons. By contrast, the dissociation of a carboxylic acid into a carboxylate anion and a proton represents a heterolytic mechanism; here both electrons are retained in one of the cleavage products (the carboxylate ion). The most important control on the bond energy is the electronegativity of the atoms bonding together. Electronegativity denominates the ability of an atom in a covalent bond to attract the shared electrons to itself (> Table 1). This will result is an asymmetric charge distribution, whose magnitude depends on the electronegativity difference between the bonding atoms. In the periodic table of the chemical elements electronegativity increases from left to right within periods and from bottom to top within groups. As a general rule, reactivity of covalent bonds increases with increasing polarity. Therefore, compounds containing exclusively nonpolar s-bonds such as saturated hydrocarbons are rather unreactive or inert (see below).

3

Saturated Hydrocarbons

3.1

n-Alkanes

The term n-alkane (5–8) refers to linear hydrocarbons with the general formula CnH2n+2. The prefix n stands for normal indicating that the molecule does not contain branches so that it represents a straight chain of carbon atoms (> Fig. 2). Each carbon atom (except the two

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Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

. Figure 2 Structures of selected saturated hydrocarbons. (5), n-butane C4H10; (6), n-pentane C5H12; (7), n-hexane C6H14; (8), n-heptane C7H16; (9), 2-methylpropane (isobutane) C4H10; (10), 2-methylbutane (isopentane) C5H12; (11), 2-methylpentane C6H14; (12), 2-methylhexane C7H16; (13), 2,2,4-trimethylpentane (isooctane) C8H18; (14), 3-methylpentane C6H14; (15), 3methylhexane C7H16; (16), 2,6,10,14-tetramethylhexadecane (phytane) C20H42; (17), cyclopentane C5H10; (18), cyclohexane C6H12; (19), decalin C10H18; (20), adamantane C10H16; (21) diamantane C14H20; (22), cholestane C27H48; (23), hopane C30H52. Isooctane (13), a branched saturated hydrocarbon which defines the 100 point on the octane rating scale, contains primary (1
), secondary (2
), tertiary (3
) and quaternary (4
) carbon atoms. Stereogenic centers are present in 3-methylhexane (15), phytane (16), cholestane (22) and hopane (23) (indicated by asterisks).

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terminal ones) is bound via s-bonds to two other carbon atoms. The two remaining free valences are occupied by hydrogen atoms, respectively; only the terminal carbon atoms have three hydrogen atoms as bonding partners. The structurally simplest organic molecule, methane (1) can be regarded as the lower end member of the homologues series of n-alkanes. There is no principal upper limit as to the chain length of n-alkanes. Polyethylene is a synthetic polymer consisting of extremely long carbon chains principally representing n-alkanes (up to 400.000 carbon atoms per molecule). A characteristic structural feature of n-alkanes is the existence of conformers. Conformers are stereoisomers which are transformed into each other by rotation about individual C–C bonds without breaking chemical bonds. The stability of different conformers depends on the bond interaction with the back lobes of orbitals on adjacent atoms, which is possible only when the atoms on adjacent atoms are staggered, and on steric repulsion. In n-alkanes the rotation barrier (= the activation energy required to transform one conformer to another) is relatively low, thus the rotatability about C–C s-bonds can be regarded as relatively unrestricted under conditions relevant to most natural environments. Methane, likely the most abundant low-molecular-weight organic compound on planet earth, is formed by both biological and geological processes. Biogenic and thermogenic methane are easily discriminated by their stable carbon isotopic compositions (for review See > Chapter 5, Vol. 1, Part 2). Isotopically light methane in deep natural gas reservoirs may be regarded as an indication for the existence of a deep subterraneous biosphere (e.g., Schoell, 1980). Conventional and unconventional (clathrate hydrates, shale gas, coal seams) gas resources represent the by far largest pool of hydrocarbons in the geosphere (e.g., > Chapter 3, Vol. 1, Part 1). Only very recently, evidence has been provided that higher natural gas hydrocarbons, i.e., ethane and propane are not exclusively formed by thermal processes but may also be produced biologically in the deep marine subsurface (Hinrichs et al., 2006). Higher n-alkanes occur as major constituents of leaf waxes of macrophytes and land plants, which can be distinguished chemotaxonomically according to the carbon number range of the homologues (e.g., Ficken et al., 2000). Due to the biosynthesis from fatty aldehydes with even-numbered carbon chains via decarbonylation (e.g., Schneider-Belhaddad and Kolattukudy, 2000), carbon numbers of biogenic alkanes typically show a significant oddover-even predominance. n-Alkanes are also the main constituents of undegraded crude oils. Here, typically no clear carbon number predominance is observed due to the unspecific formation via thermally controlled reactions. It is important to note that the majority of the global oil reserves is more or less significantly biodegraded and thus may lack n-alkanes.

3.2

Branched Alkanes

For hydrocarbons represented by the general formula CnH2n+2 more than one constitutional isomer is possible if n4. In these structural isomers the atoms are connected in different ways, thus interconversion is not possible without breaking chemical bonds. These alkanes do not possess straight chains of carbon atoms and therefore, in contrast to the normal alkanes, are termed branched alkanes. The number of possible constitutional isomers increases exponentially with increasing number of carbon atoms in the molecule. In general, the higher the degree of substitution is the more will the molecule have a spherical rather than a rod-like shape; this has a significant influence on the physical properties of isomers with implications for features such as bioavailability.

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Carbon atoms in these molecules are classified according to the number of 1, 2, 3, or 4 other carbon atoms to which they are connected as primary, secondary, tertiary, and quaternary carbon atoms, respectively, as illustrated for isooctane (13) in > Fig. 2. Tertiary and quaternary carbon atoms which are connected to four different substituents are termed chiral or stereogenic centers. Molecules containing chiral carbon atoms may exist as configurational isomers (or stereoisomers) which cannot be converted into each other without breaking of chemical bonds, although the connectivity of the atoms is identical. This class of isomers is subdivided into enantiomers, which are nonsuperimposable mirror-images of each other, and diastereomers, which are not. A structurally simple example of the former is 3-methylhexane (15), a common constituent of fossil fuels, which exists as two enantiomers, while 2,6,10,14tetramethylhexadecane (phytane) (16), another ubiquitous constituent of fossil fuels, contains three stereogenic centers and therefore exists as eight different stereoisomers (four pairs of diastereomeric enantiomers). In general diastereomers possess different physical properties, while enantiomers do not. However, in biological systems the two enantiomers of a molecule may behave different if chiral components (e.g., enzymes) are involved in a process. In fossil fuels some branched alkanes, e.g., isobutane (9), isopentane (10), 2-methylpentane (11), 3-methylpentane (14), 2-methylhexane (12), 3-methylhexane (15) etc., are relatively abundant in the molecular range up to approximately C10H22. With increasing molecular weight, individual isomers become less pronounced in comparison to the prevailing n-alkanes, however, the mixtures become more complex. It appears that biodegradability in general decreases with increasing degree of branching which is often observed as the relative enrichment of an unresolved complex mixture (UCM, also called the ‘‘hump’’) during biodegradation of crude oil and related petroleum products. The lower biodegradability of highly branched alkanes might be related to the low natural abundance of individual isomers having not favored the evolution of appropriate biodegradation pathways.

3.3

Cycloalkanes

Formally, cycloalkanes are generated by (homolytic) removal of two hydrogen atoms from two different carbon atoms in n-alkanes or branched alkanes; formation of a s-bond between these carbon atoms (which have to be interrupted by at least one carbon atom) will then result in a cycloalkane. The minimum number of carbon atoms in a cyclic hydrocarbon is three (cyclopropane derivatives) while there is no principal upper limit as to the ring size. Cycloalkanes are also called naphthenes, a term which particularly refers to a petroleum-related origin. Cycloalkanes with alkyl substituents may be called alkylcycloalkanes or cycloalkylalkanes. The structural diversity becomes even greater if the various types of polycyclic hydrocarbons are taken into account. Among these, annulated structures which formally are built up from side-on condensed cyclic segments are most prominent in naturally occurring hydrocarbon assemblages of fossil fuels. A simple example is decalin (19) which consists of two annulated cyclohexane rings. Cycloalkanes can be represented by the general formula CnH2(n+1-r) where n is the number of carbon atoms and r the number of rings in the molecule. Ring carbon atoms in cycloalkanes, as in n-alkanes and branched alkanes, are sp3-hybridized and therefore ideally should have tetrahedral bond angles of 109.5
. However, depending on the ring size, the actual structure will deviate more or less from a tetrahedral arrangement which will result in more or less pronounced ring strain. Cyclohexane (18) in the chair conformation allows almost ideal tetrahedral angles, therefore ring strain is

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

1

negligible. Similarly, no strong deviation from the tetrahedron will occur for the C–C–C bond angles in cyclopentane (17). Cyclopropane with C–C–C bond angles of 60
and, to a lesser extent, cyclobutane with C–C–C bond angles of 90
deviate most from the tetrahedral angle. They therefore are highly strained and significantly less stable than larger rings. The strain of rings with more than 6 carbon atoms varies irregularly but in general decreases with increasing ring size and is negligible in larger ring systems. In cyclohexane, the two hydrogen atoms attached to each carbon atom are chemically not equivalent. The torsional strain is the lowest if the molecule adopts the so-called chair conformation in which six of the twelve hydrogen atoms are in the plane of the ring (equatorial) while the other six are perpendicular to it (axial). Ring inversion leads to the interchange of equatorially and axially attached substituents. For spatial reasons, axial substituents interact more strongly with each other than equatorial substituents. Therefore, substituted cyclohexane conformers will generally be more stable if more of the larger substituents are in the equatorial position. The natural occurrence of rings of different size reflects their different stability. Cyclohexane and, to a lesser extent, cyclopentane moieties are by far predominating in natural products and naphthenic petroleum constituents. A broad variety of lipids in eukaryotes and prokaryotes, such as steroids, hopanoids, and other triterpenoids possess carbon skeletons which are based on annulated cyclohexane and cyclopentane rings. These important constituents of biomass are a relevant source of the structurally diverse mixtures of naphthenes found in fossil fuels. During diagenetic and catagenetic transformation of sedimentary organic matter, biogenic lipids loose functional groups and structural elements such as C–C double bonds (see below) but usually retain the original carbon skeleton. In geochemistry, hydrocarbons such as phytane (16), cholestane (22) and hopane (23), which can be regarded as chemical fossils, are called biomarkers as their carbon skeletons can directly be related to those of the respective biological precursors (for a detailed introduction to biomarkers see Peters et al., 2005). Cyclopropyl moieties occur, for example, in certain fatty acids and steroid derivatives. They are also a structural component of pyrethrins such chrysanthemic acid, a natural insecticide from Chrysanthemum cinerariaefolium and C. coccineum. Ladderane lipids produced by Anamox bacteria are an interesting example of natural products containing annulated cyclobutane rings (Sinninghe Damste´ et al., 2002). Such three- and four-membered rings normally will not survive diagenetic and catagenetic transformation of biogenic organic compounds deposited in the geosphere. Therefore, cyclopropane and cyclobutane derivatives are not relevant as constituents of fossil fuels. Likewise, there is only very limited evidence that larger rings with more than six carbon atoms play any significant role. If the fusion occurs across a sequence of atoms rather than at two mutually bonded atoms, bi- and polycyclic hydrocarbons may form bridged structures. a-Pinene (2,6,6-trimethyl [3.1.1]hept-2-ene) (34), a widely distributed constituent of plants and in particular of conifer resins, represents a typical example of a bridged hydrocarbon (additionally containing a double bond; see below). In general, bridged hydrocarbons are of subordinate relevance as constituents of fossil fuels. However, the so-called diamondoids are a class of petroleum constituents with bridged structures which apparently are generated at higher levels of thermal maturity (possibly from annulated hydrocarbons such as steranes or hopanes) (Dahl et al., 1999). These cagelike structures (adamantane (20), diamantane (21), triamantane etc.) can be regarded as representing the building blocks of diamonds (in contrast to polycyclic aromatic hydrocarbons which represent the building blocks of graphite). Polymantanes containing

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Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

up to 11 diamond-crystal cages (undecamantane) have been isolated from natural gas condensates (Dahl et al., 2003a,b). These constituents of fossil fuels appear to be highly resistant to biodegradation (Grice et al., 2000) and therefore may represent major constituents of severely altered crude oils.

3.4

Reactions of Saturated Hydrocarbons

Saturated hydrocarbons (n-alkanes, branched alkanes, and cycloalkanes) contain exclusively s-bonds but no (polar) functional groups. The difference in electronegativity of carbon (2.55) and hydrogen (2.20) is small, therefore, C–H bonds are rather nonpolar. As a consequence, saturated hydrocarbons are quite unreactive or inert. The term paraffin in fact describes this lack of affinity in chemical reactions. Efficient, specific, and selective methods for C–H bond activation in nonreactive substrates can be regarded as one of the great challenges in synthetic organic chemistry. In this perspective, any progress in understanding the biochemical (enzymatic) mechanisms of C–H bond activation – as an integral component of any hydrocarbon oxidation pathway – can potentially contribute to an advanced use of fossil fuel hydrocarbons beyond combustion. Due to the lack of polar bonds in saturated hydrocarbons, reactions with polar species via heterolytic mechanisms are relatively unimportant. In fact, the main reactions of the saturated hydrocarbons proceed via (free) radical species (= atomic or molecular species with unpaired electrons). Alkyl radicals may be generated from saturated hydrocarbons by a homolytic cleavage of a C–H or a C–C bond. This process requires significant amounts of energy, known as the homolytic bond dissociation energy (see > Table 2 for homolytic C–H bond dissociation energies of saturated hydrocarbons). The amount of energy required is related to

. Table 2 Homolytic C-H bond dissociation energies of saturated hydrocarbons (data from McMillen and Golden, 1982) Hydrocarbon

Product radical

kJ mol1

Cyclopropane

Cyclopropyl-

445

Methane

Methyl-

440

2,2-Dimethylpropane

Neopentyl-

419

Ethane

Ethyl-

411

Propane

Propyl-

410

Methylcyclopropane

Cyclopropylmethyl-

408

Cyclobutane

Cyclobutyl-

404

n-Butane

sec-Butyl-

400

Cyclohexane

Cyclohexyl-

400

Propane

Isopropyl-

398

Cyclopentane

Cyclopentyl-

396

Isobutane

tert-Butyl-

390

Cycloheptane

Cycloheptyl-

387

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

1

the stability of the formed radical which increases in the order primary < secondary < tertiary. In other words, the homolytic cleavage of a terminal C–H bond in alkanes requires more energy than the homolytic cleavage of a subterminal C–H bond. This is a possible reason for the observation that n-alkane activation in anaerobic bacteria (via radical addition to the double bond of fumarate) takes place at the subterminal but not at the terminal carbon atom, despite the unfavorable fact that this introduces an additional branch into the initial activation product (Rabus et al., 2001). Radicals may undergo a number of reaction types, including substitution, addition to double bonds, intramolecular rearrangement and fragmentation. Radical halogenation is an important process in synthetic chemistry, in which haloalkanes are formed from alkanes and molecular halogen by substitution of a hydrogen atom by a halogen atom via radical intermediates. In the production of synthetic polymers (e.g., of polyethylene from ethene) the multiple steps of chain elongation can proceed via radical addition to the double bond of the respective unsaturated monomers. Radical addition (to fumarate) plays a crucial role in the activation of many hydrocarbons in anaerobic metabolism. The involved alkyl and arylalkyl radicals are formed from their parent hydrocarbon via homolytic cleavage of a C–H bond by glycyl radical enzymes (e.g., Buckel and Golding, 2006; Heider, 2007). Combustion denominates the complete oxidation of saturated hydrocarbons to carbon dioxide and water with oxygen being the other reactant. The overall process is a complex interplay of numerous types of chemical reactions in which diverse radical species are key players. Oxygen itself is a diradical, which forms various reactive intermediates such as hydroperoxide or hydroxyl radicals. Again homolytic cleavage yielding alkyl or aryl radicals is a key step in the activation of the inert hydrocarbons to be converted to functionalized (oxygenated) species that can be further oxidized in subsequent reactions. Likewise, highly reactive oxygen species play a crucial role as cosubstrates in the activation of hydrocarbons for aerobic biodegradation (See > Chapter 14, Vol. 2, Part 4). Radical reactions are also of major importance in the formation of fossil fuels, especially natural gas and crude oil. It is generally accepted that these petroleum fluids are formed via thermal breakdown of a geomacromolecule termed kerogen (see below) during deep burial of biogenic organic matter in sedimentary basins. As temperatures increase with burial depth, pyrolytic reactions, i.e., radical reactions will lead to the fragmentation of larger structural moieties and the formation of low-molecular-weight organic compounds, predominantly hydrocarbons. Depending on the temperature-pressure-regime and the geological history, secondary cracking through radical reactions leads to further processing of the original petroleum fluids, e.g., oil-to-gas cracking is a significant process in many petroleum systems (Schenk et al., 1997). There are controversial discussions as to which extent these processes are controlled by thermal or catalytic mechanisms (e.g., Mango, 2000). Similar reactions are employed in the industrial reformation of crude oil.

4

Unsaturated Hydrocarbons

4.1

Alkenes and Alkines

Unsaturated hydrocarbons are molecules that contain at least one C–C double bond or one C–C triple bond. These types of compounds are termed alkenes (resp. cycloalkenes) or olefins (resp. cycloolefins), if the structure contains one or more double bonds, and alkines

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Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

. Figure 3 Structures of cis-but-2-ene (24) and trans-but-2-ene (25) C4H8.

(resp. cycloalkines), if it contains one or more triple bonds. Unsaturated hydrocarbons can be represented by the general formula CnH2(n+1-r-d-2t) where n is the number of carbon atoms, r the number of rings, d the number of C–C double bonds and t the number of C–C triple bonds in the molecule. The simplest alkene is ethene (3) (also ethylene), the simplest alkine is ethine (4) (also acetylene) (> Fig. 1). The term conjugated double bond denominates two or more double bonds in a molecule which are not separated by CH2 groups or other structural moieties, i.e., alternating double and single bonds. This is a significant structural element of certain naturally occurring hydrocarbons such as the carotenes lycopene (39), b-carotene (40), or isorenieratene (58). As it requires significant amount of energy to break a C–C p-bond, free rotation about C–C double bonds is essentially impossible. (This is in contrast to the saturated hydrocarbons where rotation about C–C s-single bonds is relatively unrestricted as pointed out.) As a consequence, asymmetrically substituted alkenes such as but-2-ene (24–25) may occur as two distinct constitutional isomers, which are classified according to the cis-/trans- or Z-/Enomenclature (> Fig. 3). No cis-/trans-isomerism can occur in alkenes in which at least one of the two carbon atoms forming the C–C double bond is connected to two identical substituents. In general, cis- and trans-isomers of a given alkene (or, more generally, of an unsaturated organic compound) exhibit different physical properties. Isomerization of cis- to trans-double bonds and vice versa is a physiologically significant process, e.g., the transformation of 11-cis-retinal to all-trans-retinal (and the recycling of the latter to the former) is an integral element of the vision cycle. Alkenes (and cycloalkenes) of great structural diversity occur as natural products in numerous living organisms (> Fig. 4). Even the simplest alkene, ethene, occurs as a biosynthetic product and acts as a hormone on various stages in the life cycle of plants. Many alkenes and cycloalkenes with one or more double bonds act as insect pheromones (Francke and Schulz, 1998). Carotenoids (both carotenes = hydrocarbons and xanthophylls = nonhydrocarbons) are pigments, that may be involved in energy transfer in photosynthetic organisms or act as antioxidants in living organisms in general, due to their system of conjugated double bonds. Moreover, C–C double bonds play an important role in many types of heteroatom-containing natural products, such as unsaturated fatty acids, steroids, and other triterpenoids etc. Examples of natural products containing C–C triple bonds are tridec-1-ene-3,5,7,9,11-pentaine, a polyine hydrocarbon isolated from Echinacea spp., (Z)-13-hexadecen-11-yn-1-yl acetate, a pheromone of processionary moths (Thaumetopoea spp.) and histrionicotoxin, a toxin of the poison dart frog Dendrobates histrionicus. Unsaturated hydrocarbons, in contrast to saturated and aromatic hydrocarbons, appear to play a minor role in fossil fuels. C–C double (and triple) bonds in most biogenic compounds are too reactive to survive the diagenetic and catagenetic transformations occurring in

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. Figure 4 Structures of selected unsaturated hydrocarbons. (26), (Z)-tricos-9-ene C23H46 (muscalure, sex pheromone of the housefly Musca domestica); (27), 2-methylbuta-1,3-diene C5H8 (isoprene); (28), 7-methyl-3-methyleneocta-1,6-diene C10H16 (myrcene, constituent of essential oils); (29), (3E,6E)-3,7,11-trimethyldodeca-1,3,6,10-tetraene C15H24 (a-farnesene, coating of apples and other fruits); (30), cyclohexene C6H10; (31), cyclohexa-1,3-diene C6H8; (32), cyclohexa-1,4-diene C6H8; (33), 1,4,5,8-tetrahydronaphthalene C10H12; (34), 2,6,6-trimethylbicyclo[3.1.1]hept-2-ene C10H16 (a-pinene, constituent of conifer resins); (35), 1-isopropyl-4-methylcyclohexa-1,3-diene C10H16 (a-terpinene, constituent of cardamom and marjoram oils); (36), 1-isopropyl-4methylcyclohexa-1,4-diene C10H16 (g-terpinene, various plant sources); (37), cycloocta-1,3,5,7tetraene C8H8; (38), (6E,10E,14E,18E)-2,6,14,19,23,27-hexamethyltetracosa-2,6,10,14,18,22hexaene C30H50 (squalene, shark liver oil); (39), lycopene C40H56; (40), b-carotene C40H56.

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geological systems over geological timescales. For example, hydrogenation processes with reduced sulfur species such as H2S (generated by microbial activity, i.e., bacterial sulfate reduction) as hydrogen donors may be responsible for the loss of double bonds (Hebting et al., 2006). This can nicely be illustrated by the diagenetic transformation of the already mentioned carotenoidal hydrocarbon isorenieratene (58) to isorenieratane (59), which may loose all conjugated double bonds on the aliphatic chain connecting the two benzene rings while the aromatic systems survive even elevated thermal stress due to their high stability (see below). Concentrations of olefins in crude oils are generally low (mostly below 1% by weight), although significantly higher concentrations have been reported in some instances. Curiale and Frolov (1998) reviewed various possible origins of olefins in crude oil. They may migrate with other soluble organic compounds directly from the source rock or get into oils through the process of migration-contamination, wherein light oils act as solvents for syndepositional olefins that occur along the migration route or within the reservoir section. Olefins in crude oils may also derive from ‘‘cold’’ radiolytic dehydrogenation of saturated hydrocarbons, introduced as a by-product of decay of uranium, thorium, and other radioactive elements among the reservoir minerals, or from pyrolysis due to thermal impact from igneous intrusions that occur close to the reservoired oil.

4.2

Reactions of Unsaturated Hydrocarbons

Due to the minor relevance of alkines as natural products or constituents of fossil fuels, only the reactivity of C–C double bonds is considered in this section. Alkenes undergo three main types of reactions, namely addition to the double bond, oxidation, and polymerization. Polymerization of unsaturated hydrocarbons occurs in some plants, i.e., formation of natural rubber and gutta-percha from isoprene (27). Polymerization of a wide range of unsaturated organic compounds (both hydrocarbons and nonhydrocarbons) via ionic or radical mechanisms is a key industrial process in the production of synthetic polymers. Due to the p-electrons, C–C double bonds are characterized by an elevated electron density and act as nucleophiles in the chemical reactions. Therefore, their most important reaction type is electrophilic addition in which the C–C doubled bond is converted into a C–C single bond by removal of the p-bond under concomitant formation of two new covalent s-bonds (> Fig. 5). In the first step a suitable electrophile, i.e., a species with an electron deficiency, typically a cation, forms a covalent bond with one of the two carbon atoms of the double bond while the positive charge is located on the other carbon atom. In the second step a neutral molecule is formed by connection of the originally formed cation to a nucleophile, i.e., a species with an electron surplus, typically an anion. A specific situation occurs if both reactants are unsymmetrical, e.g., two different products are possible from the addition of HCl to propene, namely 1- and 2-chloropropane. In such cases the Markownikow rule helps to predict the expected product distribution. According to the Markownikow rule, the addition of the electrophile in the first step of the reaction occurs in a mode that the more stable carbenium ion (= trivalent carbocation) is formed preferentially. The stability of carbenium ions depends mainly on the inductive effects of the substituents which may either stabilize or destabilize the positive charge. As a general rule of thumb, in alkenes the electrophile will be connected to the carbon atom of the double with the higher number of hydrogen atoms because the stability of the formed carbenium ions decreases in the order tertiary > secondary > primary due to the positive inductive effects of the alkyl groups.

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. Figure 5 General mechanism of electrophilic addition to C–C double bonds. Typical reactants X–Y that may be added to C–C double bonds in electrophilic addition reactions are molecular hydrogen (H–H), molecular halogen (F–F, Cl–Cl, Br–Br), hydrogen halide (H–Cl, H–Br, H–I) and water (H–OH).

Many electrophilic additions are reversible, i.e., alkenes (or unsaturated organic compounds) may be formed by suitable elimination reactions. Besides combustion, that completely oxidizes unsaturated hydrocarbons (in analogy to saturated hydrocarbons) to carbon dioxide and water, various reactions with different types of oxygenating reagents are known to yield specific products. Catalytic oxidation with oxygen or the reaction with percarboxylic acids yields epoxides which can be ring-opened to vicinal trans-diols by acid-catalyzed hydrolysis. The reaction of alkenes with osmium tetroxide on the other hand yields vicinal cis-diols. Ozonolysis, i.e., the reaction of alkenes with ozone, produces either ketones or aldehydes upon reductive work-up or ketones/carboxylic acids upon oxidative work-up depending on the structure of the alkene. In the case of cycloalkenes with a double bond in the ring system, diketones, dialdehydes, or oxoaldehydes are formed. It should be noted that these reactions of double bonds do not form new C–C bonds and therefore are not directly useful for (bio)synthesis of more complex carbon skeletons. However, they are very suitable for introducing functional groups. Thus, most of the synthetic reactions have enzymatic analogues which play important roles in many metabolic pathways. Hydrogenation of C–C double bonds is a step in the biosynthesis of fatty acids. Hydratation of C–C double bonds occurs in the TCA cycle (transformation of cis-aconitate to isocitrate and of fumarate to malate) and in b-oxidation of fatty acids. Various examples illustrate the relevance of oxidation of C–C double bonds for activation and further metabolism of alkenes and aromatic hydrocarbons. Alkene monooxygenase catalyses the transformation of propene to 1,2-epoxypropane, the first step in the aerobic metabolism of this hydrocarbon. Epoxidation of C–C double bonds plays a key role in biosynthetic pathways, e.g., squalene-2,3-epoxide formed from the hydrocarbon squalene (38) is a central intermediate in biosynthesis of steroids and triterpenoids. Similar monooxygenase-catalyzed reactions may also be involved in the aerobic transformation/activation of aromatic hydrocarbons. Such transformations formally are epoxidations of one specific double bond in a ‘‘cyclohexa-1,3,5-triene’’ moiety (rather than a reaction of an aromatic system) although the reaction has to overcome the aromatic resonance stabilization (see below). The resulting arene oxides may be rearranged to phenols (NIH-shift) or attacked by nucleophiles; an example of the latter is the stereospecific ring opening of arene oxides to trans-cyclohexa-3,5-diene-1,2-diol moieties. Likewise, the activation of aromatic hydrocarbons by dioxygenases resembles the synthesis of vicinal cis-diols from alkenes using osmium tetroxide. This ring hydroxylating dioxygenation is catalysed by non-heme iron monooxygenase and results in formation of a cis-cyclohexa-3,5-diene-1,2-diol moiety from

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a ‘‘cyclohexa-1,3,5-triene’’ moiety. Intra- and extradiol (ortho- and meta-)ring-cleavage in the further aerobic metabolism of catechols oxidize double bonds in a way which to a certain extent is similar to the reaction of alkenes with ozone.

5

Aromatic Hydrocarbons

5.1

Aromaticity

Aromaticity denominates a chemical property which is found in certain (but not all) cyclic molecules containing conjugated double bonds. Due to resonance stabilization, aromatic compounds are more stable than would be expected from the conjugation of the double bonds alone. The basic example of an aromatic hydrocarbon is benzene (41) (‘‘1,3,5-cyclohexatriene’’), for which two indistinguishable resonance structures exist (> Fig. 6). In benzene all six carbon atoms forming the six-membered ring are sp2 hybridized, therefore all C–C–C bond angles must be 120
, which is only possible if all carbon atoms lie in the same plane. The six unhybridized p-orbitals which are out of the plane of the atoms can freely overlap and are thought to form a cyclic molecular orbital above and below the ring plane in which the six p-electrons are delocalized. As a consequence, all six C–C bonds in the benzene ring have the same length (140 pm). In general, aromatic compounds are planar cyclic molecules with a fully conjugated double bond system which obeys the Hu¨ckel rule, i.e., it must contain (4n+2) delocalized p-electrons (n = 0, 1, 2, 3 and so on). Accordingly, cycloocta-1,3,5,7-tetraene (57), a molecule with 8 p-electrons which does exist, is not aromatic and also not planar. Any (bio) chemical reaction leading to a breakdown of the aromatic system has to overcome the resonance stabilization (151 kJ mol1 for benzene).

. Figure 6 Resonance structures of benzene (resonance between two structures is indicated by a double arrow). Aromaticity in benzene rings may also be depicted by an inner circle. (Note: Inner circles often are used in a misleading way to depict aromaticity in PAHs. Each circle represents 6 delocalized p-electrons, thus in the case of for example naphthalene two inner circles would represent 12 delocalized p-electrons, although naphthalene has only 10 delocalized p-electrons; a system with 12 delocalized p-electrons would even not obey the Hu¨ckel rule. This is best avoided if one of the possible resonance structures with its alternating double and single bonds is depicted.) Monosubstituted benzene derivatives posses three distinguishable hydrogen atoms on the aromatic ring, two in ortho- (o-; 1,2-) and meta- (m-; 1,3-) and one in para- (p-; 1,4-) position to the substituent, respectively.

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

5.2

1

Benzene Derivatives

All six hydrogen atoms in benzene, which are chemically equivalent, may be substituted by alkyl and aryl groups resulting in two principal classes of aromatic hydrocarbons, alkylbenzenes and polyphenyls. Mixed types are also possible. Three different isomers of disubstituted benzene derivatives are possible which are classified according to their substitution pattern as ortho-, meta-, or para-isomers (> Fig. 6). Environmentally the most significant are benzene (41), toluene (42), ethylbenzene (43) and the three xylene isomers (44–46) (BTEX), which occur in relatively high amounts in fossil fuels, due to their physico-chemical properties (see below) are rather bioavailable and have significant health effects. Fossil fuels contain complex mixtures of alkylbenzenes. Linear alkylbenzenes are produced industrially as intermediates in the fabrication of tensides. Biphenyl (48) is the structurally simplest representative among the polyphenyls. Especially ortho-substituted biphenyls may exhibit restricted rotatability about the C–C single bond between the two aromatic rings which can result in atropisomers in which the individual C2-isomers are optically stable. More complex polyphenyls such as o-, m-, and p-terphenyl (49–51) may occur in small amounts in fossil fuels (Marynowski et al., 2001) (> Fig. 7).

5.3

Polycyclic Aromatic Hydrocarbons

Polycyclic aromatic hydrocarbons (PAHs) are fused aromatic hydrocarbons consisting of two or more aromatic rings. The structurally simplest representative of this class of compounds is naphthalene (52). The number of condensed aromatic rings is essentially unlimited; larger PAHs can be regarded as structural subunits of graphite. Main environmental sources of PAHs are fossil fuels and incomplete combustion of organic materials. Thermogenic (origin from fossil fuels) and pyrogenic PAHs typically can be distinguished by the relative amounts of alkyl substituted derivatives versus the parent (unsubstituted) carbon skeleton which are high in petroleum-related products and low in combustion-derived products. More specific PAH ratios are used to distinguish pyrogenic or petrogenic PAHs (Yunker et al., 2002). Certain PAHs are carcinogenic, mutagenic, and/or teratogenic. While all hydrogen atoms are chemically equivalent in benzene (see above) this is typically not the case in PAHs, with few exceptions such as in coronene (57). Naphthalene contains two sets of four chemically equivalent hydrogen atoms, which are classified as a- and b-positions. Therefore two isomers of monosubstituted naphthalene derivatives exist, e.g., 1- and 2-methylnaphthalene (53–54). Anthracene (55), and phenanthrene (56) have three and five chemically nonequivalent hydrogen atoms and may thus form the corresponding number of monosubstituted derivatives, respectively. Chemically nonequivalent carbon and hydrogen atoms may behave different in (bio) chemical reactions which may result in certain regioselectivities. Furthermore, the degree of aromaticity may be different for each ring segment; e.g., in phenanthrene the central ring is less aromatic and therefore more reactive than the outer rings according to Clar’s rule (Portella et al., 2005; Randic, 2003). Based on an operational definition, the term ‘‘aromatic hydrocarbon’’ is often used for certain heterocyclic aromatic compound types such as dibenzofurans, dibenzothiophenes etc. in environmental and petroleum geochemistry. As these compounds contain heteroatoms they do not represent hydrocarbons sensu stricto. Therefore, these compound classes will be discussed in the appropriate parts of section 6.

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. Figure 7 Structures of selected aromatic hydrocarbons. (41), benzene C6H6; (42), toluene, C7H8; (43), ethylbenzene C8H10; (44), o-xylene C8H10; (45), m-xylene C8H10; (46), p-xylene C8H10; (47), 1-isopropyl-4-methylbenzene C10H14 (p-cymene, constituent of essential oils); (48), biphenyl C12H10; (49), o-terphenyl C18H14; (50), m-terphenyl C18H14; (51), p-terphenyl C18H14; (52), naphthalene C10H8; (53), 1-methylnaphthalene C11H10; (64), 2-methylnaphthalene C11H10; (55), anthracene C14H10; (56), phenanthrene C14H10; (57), coronene C24H12; (58), isorenieratene C40H48 (carotenoid of green sulfur bacteria Chlorobiaceae); (59), isorenieratane C40H66 (diagenetic product of isorenieratene).

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

5.4

1

Reactions of Aromatic Hydrocarbons

The most important reaction of aromatic hydrocarbons is electrophilic aromatic substitution, where the hydrocarbon reacts as a nucleophile due to the high electron density on the aromatic ring. In electrophilic aromatic substitution one hydrogen atom of the aromatic hydrocarbon is substituted by a functional group (> Fig. 8). The first step in the reaction mechanism is the formation of a p-complex through the interaction of an electrophile with the p-electron system. The p-complex is then converted to a s-complex in which the s-bond between the electrophile and a carbon atom of the aromatic ring already exists. The reaction is completed by removal as a proton of the hydrogen that is substituted. Classical electrophilic aromatic substitutions are halogenation, nitration, sulfonation, and Friedel-Crafts acylation and alkylation (> Fig. 8). It is important to note that the substituents in aromatic molecules have a significant effect on the course of electrophilic aromatic substitutions in a twofold manner. First, they may either activate or deactivate the aromatic system for subsequent reactions (> Fig. 8,

. Figure 8 (a) General mechanism of electrophilic aromatic substitution. (b) Selected examples. (c) The activating or deactivating influence of substituents on the reactivity compared to that of benzene is illustrated for nitration of different benzene derivatives.

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. Table 3 Classification of substituents with respect to their effects on electrophilic aromatic substitution o-/p-Directing Activating

m-Directing

-NH2, -NHR, -NR2 Amino-OH, -OR

Hydroxy-, Alkoxy-

-NHCOR

Acylamino-

-R

Alkyl-

Deactivating -F, -Cl, -Br, -I

Halogen-

-COR, CO2H

Acyl-, Carboxy-

-CONH2, -CO2R Carboxamido-, Carboalkoxy-

> Table

-SO3H

Sulfonic acid

-CN

Cyano-

-NO2

Nitro-

3). The ability of substituents to activate or deactivate an aromatic compound with respect to an electrophilic attack depends on their ability to increase or decrease the electron density on the aromatic ring by inductive or resonance effects. Benzene derivatives containing deactivating substituents such as chlorobenzene or nitrobenzene react significantly slower in electrophilic substitutions than benzene while derivatives containing activating substituents such as toluene or phenol react significantly faster. Therefore, the introduction of oxygen substituents is an important mechanism of biological activation of aromatic hydrocarbons for further metabolism. Secondly, substituents have a directing influence on the regioselectivity of subsequent reactions (> Table 3). Ortho-/para-directors typically are substituents with unshared electron pairs such as the hydroxyl group in phenol which support the attack of electrophiles in the o- and p-position by resonance stabilization of the intermediate s-complexes. Sterical effects are most important for observed deviations from the statistical distribution (2:1) of o- and p-products. Meta-directors such as the nitro group destabilize the s-complexes formed by attack of an electrophile in o- and p-position due to resonance structures with positive (partial) charges on adjacent atoms; therefore, the reaction in meta position is favored. Aromatic molecules may also react with nucleophiles, however, such reactions are not typical for aromatic hydrocarbons, i.e., the nucleophilic substitution of hydrogen in aromatic compounds is uncommon. Important mechanisms of reactions with nucleophiles are the substitution of suitable leaving groups such as chloride, bromide, or sulfite via an additionelimination mechanism (similar to electrophilic aromatic substitution) or the attack of the nucleophile on a free aryl cation that has been generated by elimination of nitrogen from an aryl diazonium salt. Some substitution reactions proceed via addition of nucleophiles to arynes (e.g., 1,2-dehydrobenzene) generated from suitable aromatic substrates by the removal of two o-substituents. Nucleophilic substitution plays a role in coupling reactions such as the Ullmann reaction used to synthesize biaryls. Homolytic cleavage of aromatic C–H bonds is energetically unfavorable (the homolytic C–H bond dissociation energy to form the phenyl radical from benzene is as high as 464 kJ mol1; McMillen and Golden, 1982). Therefore, reactions of aromatic molecules via free aryl radical intermediates are not very common. An interesting reaction involving aryl radicals is the hydroxylation of benzene to phenol by Fenton’s reagent. Fenton’s reagent, a solution of

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hydrogen peroxide and an iron catalyst which generates hydroxyl and hydroperoxide radicals as reactive species, is used to oxidize contaminants and waste water. Aromatic hydrocarbons may be catalytically hydrogenated to the corresponding cycloalkanes at high temperature and pressure. Reactions of aromatic hydrocarbons leading to a partial reduction of the aromatic ring are rare because of the high resonance energy. In fact, cyclohexa-1,3-diene (31) and cyclohexa-1,4-diene (32) are excellent hydrogen donors in transfer hydrogenation as they readily oxidize to benzene. The reduction of benzene (41) to cyclohexa-1,4-diene (32) with sodium and an alcohol in liquid ammonia is known as the Birch reduction. Likewise, naphthalene (52) may be reduced under these conditions to 1,4,5,8tetrahydronaphthalene (33). The Birch reaction proceeds via alternating single electron- and proton-transfer steps to the aromatic ring. It is assumed that the enzymatic reduction of benzoyl-CoA, a key step in anaerobic metabolism of aromatic compounds proceeds via a Birch-like mechanism, although the product is a conjugated 1,3-diene and not a nonconjugated 1,4-diene (Boll et al., 2002). It is worth mentioning that hydrocarbons with a cyclohexa1,3- or -1,4-diene moiety occur as natural products, e.g., a- and g-terpinene (35–36), however, biosynthesis does not proceed via reduction of the corresponding aromatic hydrocarbon p-cymene (47) which is a common constituent of essential oils.

6

Functionalized Organic Compounds

Functionalization of aliphatic and aromatic hydrocarbons-formally by insertion of heteroatoms into carbon–carbon or carbon–hydrogen bonds-expands dramatically the structural diversity of organic compounds and alters essentially their chemical and physico-chemical properties. Most important heteroatoms in organic chemistry are oxygen, nitrogen, and sulfur, followed by the group of halogens and to a minor extent by phosphorus. The insertion of functional groups is not restricted to monofunctionalization or to one heteroatom involved. Noteworthy, certain multifunctionalized molecules can undergo intramolecular coupling reactions leading to heterocyclic systems, whereas intermolecular interactions result in coupling of two (or more) molecules.

6.1

Halogens

Halogen atoms are attached to aliphatic as well as aromatic moieties. From a formal point of view halogen atoms substitute hydrogen atoms in C–H bonds resulting in alkyl and aryl halides. Chemical reactions leading to halogenated organic compounds include radical, nucleophilic, or electrophilic substitutions of hydrogen atoms in hydrocarbons or functional groups in functionalized compounds (primarily alcohols). Predominantly chlorine and bromine are organically bound halogens, whereas fluorinated and iodinated compounds are less common, especially in natural products (prominent exceptions are sodium fluoroacetate, an antiherbivore poison in various plants, or the iodine containing thyroid hormones, thyroxine and triiodothyronine). However, fluorine-containing compounds are still important xenobiotics with partially high environmental impact, e.g., chlorofluorocarbons (CFC) or perfluorinated detergents (PFAS, PFCA). Even for iodine-organic compounds some very special applications are known, e.g., the iodine-containing X-ray contrast medium iopromide, leading also to an environmental occurrence of such compounds.

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. Table 4 Structural diversity of halogenated benzenes

Structural element

Variability

Resulting number of congeners

Degree of halogenation

x = 1 to 6

6

Substitution pattern

Mixed halogenation (example restricted to bromine and chlorine)

Cl1: 1

Cl1; Br1: 2

Cl2: 3

Cl2; Br2; BrCl: 9

Cl3: 3

Cl3; Br3; BrCl2; Br2Cl: 17

Cl4: 3

Cl4; : Br4; BrCl3; Br2Cl2; Br3Cl: 40

Cl5: 1

Cl5; Br5, BrCl4; Br2Cl3; Br3Cl2; Br4Cl: 20

Cl6: 1

Cl6; Br6; BrCl5; Br2Cl4; Br3Cl3; Br4Cl2; Br5Cl: 13

12

101

The high structural diversity of halogenated organic compounds is based on three independent factors. First, the degree of halogenation may cover a wide range from monohalogenated to perhalogenated compounds. Secondly, the location of halogen substituents at different carbon atoms of aliphatic or aromatic moieties leads to numerous substitutional isomers. Finally, mixed halogenation strongly expands the compositional variability. Thus halogenated compounds with a given carbon skeleton exist as large sets of so-called congeners, as exemplified for the chlorinated/brominated benzenes in > Table 4. Numerous congener groups of xenobiotic halogenated organic compounds represent important environmental contaminants featuring the described structural diversity. Polychlorinated dibenzo-p-dioxins and dibenzofurans (usually summarized as ‘dioxins’) exhibit specific congener patterns allowing to differentiate their typical emission sources (e.g., Fiedler, 1996). Also polychlorinated biphenyls (PCBs) or polybrominated diphenyl ethers (widespread used flame retardants) appear with characteristic patterns of congeners (maximum number of congeners: 209, respectively) (Ballschmiter et al., 1987; de Boer et al., 2000). Over decades it has been assumed, that halogenated compounds can be found only infrequently in living organisms. However, over the last 20 years the information on natural chlorinated and brominated compounds increased dramatically and indicated a high diversity of halogenated natural products in fungi, algae, terrestrial plants, mammals, and further biota (Gribble, 1994, 2000; Ballschmiter, 2003). The molecular structures range from simple haloalkanes to complex and highly functionalized compounds as illustrated in > Fig. 9. It is also shown, that a high degree of halogenation is not restricted to xenobiotic compounds but can be also observed in natural products.

6.2

Oxygen and Sulfur

Since chalcogens are divalent elements they are able to form either single or double bonds with carbon atoms. Inserting oxygen or sulfur into C–H bonds leads to alcohols in the case of

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. Figure 9 Halogenated organic compounds naturally produced by living organisms (according to Gribble, 1994, 2000). (60), bromomethane CH3Br (marine algae); (61), 1-bromo-2-iodoethane C2H4BrI (marine algae); (62), 1,1-dibromo-4,4-dichlorobut-3-en-2-one C4H2Br2Cl2O (red alga Asparagopsis taxiformis); (63), (1S,2R)-3,4-dibromo-5-methylenecyclopent-3-ene-1,2-diol C6H6Br2O2; (64), 2,4,5-tribromo-3-(2,3,5-tribromo-1H-indol-1-yl)-1H-indole C16H6Br6N2 (cyanobacterium Rivularia firma); (65), 3,4,5-trichloro-2-(2,3-dichloro-1H-pyrrol-1-yl)phenol C10H4Cl5NO (bacterium Streptomyces sp.); (66), 2-amino-3-(3-bromo-5-chloro-4-hydroxyphenyl)propanoic acid C9H9BrClNO3 (mollusc Buccinum undatum).

aliphatic or phenols and in the case of aromatic moieties as well as the sulfur-analog thiols and thiophenols. The corresponding oxygen-containing functional group is called hydroxyl group, whereas for S–H groups the use of a prefix in nomenclature is unusual. Since sulfur has a lower electronegativity than oxygen (> Table 1), a higher polarity and bond strength of the CO single bond and, correspondingly, different bond length of C-O (ca. 143 pm) and C-S (ca. 180 pm) single bonds as well as differing reactivity are evident. Based on their high polarity hydroxyl groups are forming strong so-called hydrogen bonds, special intermolecular forces between partially positively charged hydrogen atoms and partially negatively charged heteroatoms (like oxygen) of neighbored molecules (see also section 8). Such strong intermolecular interactions are not realized between thiols, but on the contrary thiols exhibit a higher acidity in comparison to alcohols as the result of lower bond strength of S–H as compared to O–H bonds. Hydroxyl groups can be attached either to aliphatic or aromatic moieties, the latter representing the phenols named according to the simplest representative of this compound class, hydroxybenzene or phenol. It is noteworthy that a direct linkage of hydroxyl groups to aromatic rings influences rigorously the acidity of the functional group. Aliphatic alcohols exhibit only a weak acidity and, consequently, strong bases (e.g., elemental alkali metals or their hydrides) are needed to generate the corresponding salts, the alcoholates. The acidity depends predominantly on the inductive effects of further substituents. Thus, perfluorination of the methyl group in ethanol (high negative inductive effect as the result of the high electronegativity of fluorine) shifts the pKa value from 16 (ethanol) to 12.5 (2,2,2-trifluoroethanol (69)). On the contrary, the pKa value of 10 of phenol indicates a relatively high acidity as the result of resonance stabilization, hence, phenols react principally as weak acids.

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However, their acidity is also influenced by inductive effects of further substituents (e.g., halogens) attached to the aromatic ring. The reactivity of aliphatic alcohols is also influenced by the structural properties of the hydroxylated carbon atoms. Three different types, namely primary (e.g., 2,2,2-trifluoroethanol (69), carbon atoms 1 and 3 in glycerol (70)), secondary (e.g., carbon atom 2 in glycerol (70)) and tertiary (e.g., tert-butanol (75)) alcohols are differentiated according to the number of 1, 2 or 3 carbon atoms to which the hydroxylated carbon atom is bonded. Due to differences in resonance stabilization the reactivity of these different alcohols varies. For example, the type of elimination reactions shifts from a more unimolecular mechanism (E1) in tertiary alcohols to a more bimolecular mechanism (E2) in primary alcohols. More than one hydroxyl group may be attached to aliphatic and aromatic moieties in diverse compound classes. In case of two neighbored hydroxyl groups the term vicinal substitution (vic-) has been established. Polyhydroxybenzene derivatives are well known metabolic intermediates and mono- or dihydroxylation of aromatic rings is a key reaction in degradation pathways of aromatic hydrocarbons. However, aliphatic polyols play a more prominent role in biochemistry. 1,2,3-Trihydroxypropane (70), named glycerol, represents the backbone of fat molecules (triglycerides), in which it is linked to three fatty acids via ester bonds, and of glyco- or phospholipids. A very high degree of hydroxylation is also realized in carbohydrates, where most of the carbon atoms carry a hydroxyl group leading to a high relative oxygen content. A further interesting example of aliphatic polyols is shikimic acid (73), an intermediate of biosynthesis of several important aromatic compounds (e.g., phenylalanine, tryptophane, gallic acid etc.). In geminal (gem-) diols two hydroxy groups are bonded to the same carbon atom. Typically, these compounds are unstable and form carbonyl compounds (see below) by elimination of water. Only very few gem-diols such as formalin (71) and chloral hydrate (72) are stable under normal conditions. Alkylated phenols are common constituents of crude oil; likewise, phenolic moieties occur widespread in biogeomacromloecules (lignin, humic substances, kerogen; see section 7). In contrast, aliphatic alcohols are less represented in geologic organic matter. However, alcohols are a fundamental compound class in the technosphere. Methanol, the simplest aliphatic alcohol, is one of the most important industrial chemicals with an annual production rate of around 30 million tons. Further specific alcohols used in plasticizers and additives are 2-ethylhexanol (74) and tert-butanol (75). An exchange of the hydrogen atom in hydroxyl groups by aliphatic or aromatic moieties leads to the compound classes of ethers. The sulfur analogues are named thioethers. Principally, alkyl/alkyl-, aryl/aryl- and alkyl/aryl ethers exist in cyclic (e.g., tetrahydrofuran, 1,4-dioxan, polychlorinated dibenzo-p-dioxins and dibenzofurans PCDD/F) or acyclic constitution (e.g., diethyl ether (76), diphenyl ether (77), anisole (78)). The smallest cyclic ether, oxirane, consists of a three-membered ring, and is highly reactive due to high ring strain; it also occurs as a functional group, the epoxy group, in larger molecules. The generation of epoxides by insertion of oxygen into C=C double bonds is an important reaction to form reactive intermediates in biochemistry. For example, epoxidation is the starting reaction for the cyclization of steroids from squalene or for the initial activation of aromatic hydrocarbons as a part of their degradation pathways. Cyclic ether moieties of higher stability (lower ring strain) occur in numerous natural products (e.g., a-tocopherol (79)) and xenobiotics (e.g., crown ethers). The chemical behavior and the physico-chemical properties differ significantly from those of alcohols. Most ethers are less reactive and, consequently, are often used as solvents in the

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chemical industry or inert additives in commercial products (e.g., methyl-tert-butyl ether MTBE (80) as anti-knocking agent in gasoline). Thioether moieties are also present in natural products (e.g., dimethyl sulfide (81), methionine (82)) as well as in xenobiotic compounds (e.g., the chemical warfare agent bis(2-chloroethyl)sulfide (83) known as mustard gas or S-Lost). An important aspect of ether units in the technosphere is polyether synthesis using ethylene oxide leading to the polymer group of polyethylene glycols or polyethylene oxides with molecular weights of up to 10.000.000 g mol1 widespread used in industrial, pharmaceutical, medicinal and personal care products, in detergents, and as additives in other polymers. The C–O linkage is not restricted to single bonds, also the C=O double bond forming the carbonyl group is an important moiety in organic compounds. The formation of C=O double bonds needs the reorganization of the sp3-hybridization of the carbon and oxygen atoms in C–O single bonds to sp2-hybrid orbitals for generating the C–O s-bond as well as the p-bond by interaction of the remaining p-orbitals. As a result the geometry of this functional group is planar. In the case of asymmetrically substituted carbonyl groups both plains (above and below the planar group) are enantiotopic. Thus, the direction of addition of a nucleophilic reaction partner to the carbon atom determines the stereochemical properties of the generated enantiomer. Carbonyl groups occur in two different compound types, the aldehydes, in case of terminal attachment, and ketones, in which the C=O double bond is located at secondary carbon atoms. The conversion of primary alcohols to aldehydes or secondary alcohols to ketones is an oxidation reaction. Ketones may also occur in cyclic structures, e.g., cyclohexanone. The need for two free valences prohibits the formation of C=O double bonds directly at aromatic carbon atoms. However, the specific cyclic structure of p-quinone (cyclohexa-2,5-diene-1,4-dione) is integrated in many natural products, especially in plant pigments or vitamins (vitamin K (84)), but is also a known moiety in abiotically formed oxidation products of PAHs (e.g., anthra-9,10-quinone (85) derived from photo-oxidation of anthracene). As a result of the high polarity of the carbonyl group, addition (as mentioned above) and condensation reactions dominate the chemical behavior of aldehydes and ketones. The primary reaction step is the attack of the carbon atom by a nucleophilic reagent. However, for several ketones and, to a minor extent, aldehydes two different reaction characteristics exist in parallel. Carbonyl groups exhibit the so-called keto-enol tautomerism, in which two different forms (tautomers) of one molecule, the carbonyl form and an unsaturated alcohol, coexist in a rapid equilibrium. The interconversion of tautomers requires the shift of s- and pbonds as well as the transfer of one hydrogen atom via the enolate ion. For most of the ketones and nearly all aldehydes the keto tautomer is energetically favorable and thus much more abundant (enol tautomer of acetone approximately 0.00025%). However, in b-oxocarbonyls an extended resonance and the additional inductive effect of the second C=O bond induce a higher stability of the enol tautomers highly influenced by the solvent (enol tautomer of ethyl acetoacetate (67) in water approx. 0.4%, in hexane approx. 46.4%). Further, the a-hydrogen atom can be easily abstracted as a proton, because of the mesomeric stabilization of the corresponding anion. This effect leads to an elevated acidity of b-oxocarbonyls. An interesting example of a ‘‘frozen’’ keto-enol tautomer is the biochemically important phosphoenolpyruvic acid (PEP) (68), which from a formal point of view is the ester of phosphoric acid and pyruvic acid in its enol form (> Fig. 10). Acetals and hemiacetals are compounds with two C–O single bonds at one carbon atom but without a double bond. These derivatives of aldehydes and ketones are formally two ether

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. Figure 10 Keto-enol-tautomerism as exemplified for ethyl acetoacetate C6H10O3 (67) whose enol-form is stabilized by an intramolecular hydrogen bond, and a ‘frozen’ example, phosphoenolpyruvic acid C3H5O6P (68).

groups or one ether and one hydroxyl group attached to one carbon atom and, therefore, represent mono- or dialkylated gem-diols. In contrast to the unstable gem-diols, acetals exhibit a higher stability, whereas hemiacetals are also of minor stability. However, the most important natural products exhibiting hemiacetal groups are carbohydrates in their cyclic form, in which one hydroxyl group has reacted intramolecularly with the keto or aldehyde group. Further intermolecular reactions of these hemiacetals with hydroxyl groups of other carbohydrate molecules form acetals and the frequent repetition of this intermolecular reaction by numerous monomers build up oligo- and polysaccharides (see section 7). A C–O single and a C=O double bond at one carbon atom are realized in carboxylic acids, the oxidation products of aldehydes, and certain of their derivatives. Abstraction of a proton from the hydroxyl group of carboxylic acids results in an anion highly stabilized by resonance which explains the high acidity of this compound class. Beside acid/base reactions the carboxy group tends to exchange the hydroxyl group by other functional moieties like amines, alkoxy groups, or halogen forming carboxylic acid derivatives (amides, esters, and acyl halides). In particular amides and esters are important compounds in biosphere and technosphere. The condensation reaction of carboxylic acids and amines forming amides is important to build up peptides and proteins from amino acids. Esters can also be formed by inorganic acids (sulfuric acid, phosphoric acid, nitric acid, etc.) and alcohols. Phosphoric acid and carbohydrate derivatives are the monomers building up the backbone of the nucleic acids (DNA and RNA, see section 7). Cyclic esters, referred to as lactones, are realized in numerous natural products. Remarkable examples are the so-called macrolides, cyclic biomolecules with rings of usually 14–16 atoms (maximum variation of ring size 6–62) frequently used as antibiotics. Cyclic amides, the so-called lactams, are of lesser importance and their structural properties are less diverse as compared to lactones. However, the core structure of penicillin consists of a four membered lactam ring, and five to seven ring lactams are useful starting materials for technical polyamide synthesis (e.g., e-caprolactam (86)). Beside sulfur analogues of alcohols and ethers, sulfur also occurs in other more specific functional groups. One special structural feature of sulfur is the possibility to form oligosulfides by insertion of sulfur atoms into S–S–, S–H– or S–C bonds of thioethers or thiols. This reaction is the result of a weak oxidation, during which the oxidation state of the sulfur atoms increases with ongoing rate of insertion. Disulfides are also formed by an oxidative coupling of

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two thiols. Since this reaction is reversible, the thiol/disulfide redox reaction system plays a particular role in the three-dimensional arrangement of various biomacromolecules (e.g., enzymes and hormones). Naturally occurring polysulfides with a very simple molecular structure are dimethyl disulfide and dimethyl trisulfide (87). Since sulfur must not fulfill the octet rule exactly, sulfur containing functional groups can further be oxidized by adding oxy and hydroxyl groups with single and double bonds to the sulfur atom. In case of thioethers, the resulting functional groups are sulfoxides and sulfones. Exchanging an alkyl or aryl substituent in sulfones by a hydroxyl group or amines leads to the compound classes of sulfonic acids and sulfonamides, which are frequently found in technical products, e.g., plasticizers, pharmaceuticals, or detergents (N-butylbenzenesulfonamide NBBS (88), linear alkylbenzenesulfonates LAS (89)). Special molecular structures, in which oxygen as well as sulfur atoms are involved in aromatic systems with sp2-hybridization, are five-membered rings containing these heteroatoms. The conjugated carbon–carbon double bonds form together with the remaining p-orbital of the heteroatom the delocalized aromatic p-system. Basic compounds of this group of so-called heterocyclic aromatic compounds are furan and thiophene. Their aromaticity is lower (lowest for furan) when compared to aromatic hydrocarbons sensu stricto, but they exhibit typical aromatic properties with respect to their reactivity, spectroscopic behavior, and thermodynamic stability. Higher ring systems are built up by fusion of further benzene moieties leading to benzofuran, dibenzofuran or benzothiophene, dibenzothiophene or benzonaphthothiophenes (90–94), respectively. Furan and thiophene derivatives are common constituents of coals, tar, and petroleum. Furthermore, one of the most prominent group of anthropogenic pollutants, the dioxins or PCDD/PCDFs, consist to a large part of chlorinated congeners of dibenzofuran (> Fig. 11).

6.3

Nitrogen

The elements of the fifth main group of the periodic table of the chemical elements, that are involved significantly in organic chemistry, are nitrogen and to a much lower extent phosphorus. The latter element appears in organic compounds dominantly as phosphoric acid coupled with organic moieties via ester bonds. Although the structural diversity of organically bound phosphorus is very limited, certain of these compounds are of essential relevance in biochemical processes. Well-known examples highlighting this importance are adenosine tri- and diphosphate (ATP, ADP), the nucleic acids (RNA, DNA), phospholipids, or phosphoglycerates. The structural diversity of nitrogen-containing organic compounds is much higher and comparable to that of oxygen- and sulfur-containing compounds. A group of simple nitrogen-containing compounds is obtained by sequential alkylation of ammonia forming the amines which, depending on the degree of alkylation, are classified as primary, secondary, and tertiary amines. Compared with the ammonium cation, quaternary alkylation results in the formation of quaternary ammonium cations and corresponding salts. The formation of aliphatic ring systems in which secondary or tertiary amines are involved are common structural moieties (e.g., pyrrolidine (95), piperidine (96), cocaine (97), morphine, strychnine). Further, the functional group of amines, the amino group, can be attached manifold to carbon backbones leading to diamines, triamines etc. such as spermidine (98) (a biogenic compound involved in cellular metabolism), in which a secondary amine is further substituted

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. Figure 11 Structures of selected oxygen and sulfur containing compounds. (69), 2,2,2-trifluoroethanol C2H3F3O; (70), glycerol C3H8O3 (1,2,3-trihydroxypropane); (71), formalin CH4O2 (methanediol); (72), chloral hydrate C2H3Cl3O2 (2,2,2-trichloroethane-1,1-diol); (73), shikimic acid C7H10O5 ((3R,4S,5R)-3,4,5-trihydroxycyclohex-1-ene-1-carboxylic acid); (74), 2-ethylhexanol C8H18O; (75), tert-butanol C4H10O; (76), diethyl ether C4H10O; (77), diphenyl ether C12H10O; (78), anisole C7H8O (methoxybenzene); (79), a-tocopherol C29H50O2; (80), methyl-tert-butyl ether MTBE C5H12O; (81), dimethyl sulfide C2H6S; (82), methionine C5H11NO2S (2-amino-4-(methylthio)butanoic acid); (83), bis(2-chloroethyl)sulfide C4H8Cl2S (S-Lost); (84), vitamin K; (85), anthra-9,10-quinone C14H8O2; (86), «-caprolactam C6H11NO (azepan-2-one); (87), dimethyl trisulfide C2H6S3 (dimethyltrisulfane); (88), N-butylbenzenesulfonamide NBBS C10H15NO2S; (89), linear alkylbenzenesulfonates LAS; (90), benzofuran C8H6O; (91), dibenzofuran C12H8O; (92), benzothiophene C8H6S; (93), dibenzothiophene C12H8S; (94), benzo[b]naphtho[2,1-d]thiophene C16H10S.

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by two amino groups forming a triamine. In analogy to the corresponding oxygen compounds (alcohols, phenols) also amines form intermolecular hydrogen bonds. The replacement of hydrogen atoms in ammonia by alkyl groups enhances the basicity by a positive inductive effect, which can partially be compensated by an opposite direction of the inductive impact or by steric hindrance. Therefore, first and second alkylation leads to a slightly elevated basicity, whereas tertiary amines exhibit similar pKb values when compared to ammonia. On the contrary, amines with aromatic substituents exhibit no distinct alkalinity as a result of the strong negative mesomeric effect of the aromatic substituent. This compound class refers to anilines denominated according to its simplest member, phenylamine or aniline (99). Amines are widespread constituents of the biosphere. Amino substitution is a basic structural feature of amino acids and enables this group of essential compounds to build up peptides and proteins by polycondensation (see section 7). Other important biogenic nitrogen compounds act as neurotransmitters (e.g., serotonin (100), dopamine (101)), or hormones (e.g., histamine (102), adrenaline (103)). Volatile aliphatic amines exhibit bad odor and represent degradation products of more complex nitrogen containing compounds. In the technosphere amines are frequently used directly or act as raw material for synthesis of dyes, in particular azo dyes (e.g., methyl orange (104)), and drugs (amphetamine (105) and derivatives). As introduced in section 6.2 amines as well as anilines readily react with carboxylic acids to form amides or anilides, respectively. In accordance with the nomenclature of substituted amines, amides are also differentiated by their degree of substitution as primary, secondary, and tertiary amides. Of particular interest is the stabilization of carbonic acid by the formation of its mono- or diamide resulting in carbamate or urea derivatives. Besides the natural occurrence these structural moieties are building up important classes of pesticides. Pesticides with carbamate or urea structure are, for example, aldicarb (106) and isoproturon (107), respectively. Noteworthy, the formation of amides is not limited to carboxylic acids but also possible with sulfonic acids (sulfonamides; see section 6.2) and phosphoric acid. Carbon–nitrogen bonds are also realized as double or triple bonds. Imines, which exhibit a C–N double bond, are of minor importance with respect to structural diversity. However, imine formation is known as an initial reaction in the nonenzymatic browning reaction between amino acids and carbohydrates, the so-called Maillard reaction. Also, the C–N triple bond, the functional group of nitriles, is of elevated significance solely in the technosphere where, for example, acetonitrile is an important solvent and the polymerization of nitriles, especially acrylnitrile, forms resistant polymers. However, Fleming (1999) reviewed various nitrile-containing natural products. From a chemical point of view nitriles are classified as carboxylic acid derivatives, since depletive hydrolysis of nitriles yields carboxylic acids. The nitrogen atom in amino groups is amenable for oxidation forming the nitro group. Principally, nitro groups can be attached to aliphatic and aromatic moieties. However, in terms of environmental occurrence only the nitro arenes are of greater importance. The nitro group exhibits an elevated relative oxygen content, which is the reason for the highly explosive properties of multinitro arene derivatives. Best known nitro containing explosives are 2,4,6trinitrotoluene (TNT) (108), tetryl (2,4,6-trinitrophenyl-N-methylnitramine (109)) and picric acid (2,4,6-trinitrophenol (110)). Nitro substitution can also be found in selected personal care products, especially as fragrances like musk xylene (111) or musk ketone and pesticides (e.g., nitrofen (112)). Nitro compounds should not be confused with organic nitrates, which represent the esters of nitric acid with alcohols forming explosives (e.g., nitroglycerin (113)) and are also used in pharmaceuticals (e.g., isosorbide mononitrate (114)).

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Nitrogen is often incorporated in aromatic systems. With respect to sulfur and oxygen, heterocyclic aromatic compounds are dominated by five-membered ring systems containing one heteroatom. However, the possibility to form heterocyclic aromatic systems is not restricted to five-membered rings or to one heteroatom. In particular, nitrogen is forming a more complex family of heterocyclic aromatic compounds including different ring sizes with one (e.g., pyrrole, pyridine, azepine cation) or more nitrogen atoms (e.g., imidazole, pyrazine, triazine). Bicyclic structures may contain nitrogen atoms in one or both ring systems (quinoline and purine). Despite their aromatic character many of these compounds exhibit weak basicity. These structural moieties appear widespread in natural products comprising alkaloids, amino acids, nucleic acids, and chlorophylls (> Table 5). Xenobiotics with nitrogen containing heteroaromatic moieties are the triazine pesticides including atrazine (115), simazine and terbuthylazine (> Fig. 12).

6.4

Specific Reactions of Functionalized Organic Compounds

Since functional groups influence the polarity in organic molecules, these moieties are primary regions of chemical reactivity. Hence, functional groups act as dominant reaction centers in molecules and control their reaction behavior. Principally, three different types of reactions, in which functionalized compounds are involved, can be differentiated, namely (1) the conversion or substitution of functional groups, (2) the linkage of two individual molecules via heteroatoms, and (3) the linkage of two molecules by C–C bonds. Furthermore, as introduced in this section a second and more common system for classification is based on the reaction mechanism comprising elimination, addition, intramolecular rearrangements, condensation, and substitution reactions and, moreover, oxidation and reduction. However, some reactions introduce functional groups in formerly nonfunctionalized molecules or delete functional groups to generate nonfunctionalized compounds. This reaction type comprises elimination and addition reactions, which are inverse reaction mechanisms. They are characterized either (1) by the abstraction of functional groups or moieties accompanied by the loss of a second moiety (mostly hydrogen atoms) leading to the generation of unsaturated hydrocarbons or (2) by the addition of reactive agents to C–C double bonds introducing new functional groups. Therefore, principal aspects of these reactions are discussed in sections 3.4 and 4.2. Reactions leading to the exchange of functional groups are predominantly substitution reactions. Two reaction types are distinguished, based on the affinity of the attacking agent, the nucleophilic and the electrophilic substitution. A third substitution mechanism in which radicals are involved is of minor importance in functional group chemistry, since a radical mechanism implies a homolytic cleavage as an initial reaction step. Normally, such homolytic cleavage is hindered in functionalized moieties because of the asymmetrically distributed electron density in functional groups and neighboring moieties. Electrophilic substitution reactions, initiated by agents with a depleted electron density attacking molecule moieties with an elevated electron density, proceed typically at aromatic systems, since the delocalized p-electron system is highly attractive for electrophiles. However, electrophilic substitutions at aromatic ring systems are often characterised by the introduction of new functional groups, and not by the exchange of existing functional moieties (see section 5.4). Nucleophilic substitution is of great relevance for fuctional groups in which more electronegative heteroatoms shift off negative electronic charge from the attached carbon atoms; in nucleophilic substitution reactions the nucleophilic agents primarily attack these carbon

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. Table 5 Selected nitrogen-containing heterocyclic aromatic systems and examples of biogenic compounds containing these moieties pyrrol

hemoglobin, chlorophyll

pyridine

nicotine

imidazole

caffeine, histidine

indole

tryptophane

quinoline

quinine

purine

adenine (DNA, ATP/ADP)

atoms. Typical nucleophiles are anions or agents with atoms exhibiting free electron pairs such as ammonium and amines, halides, hydroxide ions, alkoxylates, and sulfur analogues. Important nucleophilic substitution reactions are, for example, the conversion of alcohols to halides, the methylation of alcohols generating methyl ethers, or the alkylation of primary amines to form secondary or tertiary derivatives. Depending on the number of reactants involved in the rate determining step of substitution reactions (one or two), a mono- and bimolecular reaction mechanism has to be distinguished. Besides kinetic effects these mechanistic differences have implications for the stereochemistry of the substitution product. In monomolecular substitutions (SN1) a triplanar ionic transition state is attacked in the second reaction step from two sides or directions leading to racemization in the case of prochiral compounds. On the contrary, a bimolecular reaction (SN2) proceeds through a fixed bipyramidal structure of the intermediate and thus results in the inversion of the stereochemistry. Note that nucleophilic substitutions are not restricted to the exchange of functional groups but are useful tools to link individual molecules via heteroatoms. For example, ether bridges are generated by the nucleophilic attack of alkoxy ions on appropriate substrates; they are key structural elements in archaeal and bacterial lipids (isoprenoidal and non-isoprenoidal glycerol diethers). The inverse reaction, in which the ether bond cleavage is typically initiated by strong Lewis acids, also ranks among nucleophilic substitutions. A specific reaction type in which solely functionalized molecules are involved is the condensation. This reaction is

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. Figure 12 Structures of selected nitrogen containing compounds. (95), pyrrolidine C4H9N; (96), piperidine C5H11N; (97), cocaine C17H21NO4; (98), spermidine C7H19N3 (N-(3-aminopropyl)butane-1,4diamine); (99), aniline C6H7N; (100), serotonin C10H12N2O (3-(2-aminoethyl)-1H-indol-5-ol); (101), dopamine C8H11NO2 (4-(2-aminoethyl)benzene-1,2-diol); (102), histamine C5H9N3

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characterized by the linkage of two functional groups leading to a larger product molecule accompanied by the release of a second small sized molecule. This second product is typically a thermodynamically highly stable compound (H2O, HCl or similar), whose generation represents the propulsion of the reaction. Basically, condensation reactions are a subcategory of substitution reactions. However, condensation reactions are important to generate linkages of two individual molecules via heteroatoms. Essential condensation reactions are performed by carboxylic groups with alcohols and amines forming esters and amides. These condensation reactions are used not only to build up dimers (e.g., aspartame) but also (in case of polyfunctionalized molecules) to form oligomers (e.g., valinomycin) or natural as well as xenobiotic polymers (e.g., proteins, suberins, nylon). Besides intermolecular reactions, intramolecular substitution and condensation occur widespread. In particular, for oxygen containing compounds, the formation of (1) cyclic ethers via nucleophilic substitution, (2) cyclic acetals and hemiacetals via addition or condensation, or (3) esters via condensation have already been mentioned in section 6.2. As a result of heteroatom functionalization, the state of oxidation at the affected carbon atoms may change. By introducing a more electronegative binding partner or more bonds to heteroatoms, the oxidation state shifts to a more positive value as the result of depleted electron density at the carbon atom. Reactions leading to increasing oxidation states are oxidation reactions and the inverse reactions are reduction reactions. Note that oxidation and reduction are not limited to carbon atoms since heteroatoms also exhibit different oxidation states. Essential oxidation reactions are the sequential transformation of primary alcohols to carboxylic acids via aldehydes as exemplified in the initial steps of the aerobic degradation pathway of citronellol forming citronellic acid. In this reaction sequence two new carbon–oxygen bonds are introduced by shifting the oxidation state of the involved carbon atom. Oxidation/reduction systems involving carbon and heteroatoms may act as sensors or indicators for environmental conditions with respect to oxygen availability. Transformation of nitro groups to the corresponding amino groups (as observed in transformation reactions of explosives (TNT) or nitro pesticides), indicates an oxygen depletion. The oxidation/reduction sequence of selected sulfur organic compounds responds sensitively to changing redox conditions, for example, in the formation of sulfoxides and sulfones from sulfides or vice versa.

(2-(1H-imidazol-4-yl)ethanamine); (103), adrenaline C9H13NO3 (4-[1-hydroxy-2-(methylamino) ethyl]benzene-1,2-diol); (104), methyl orange C14H14N3O3SNa (4-{(E)-[4-(dimethylamino)phenyl] diazenyl}-2-methylbenzenesulfonic acid); (105), amphetamine C9H13N (1-methyl-2phenylethylamine); (106), aldicarb C7H14N2O2S ((1E)-2-methyl-2-(methylthio)propanal O[(methylamino)carbonyl]oxime); (107), isoproturon C12H18N2O (N0 -(4-isopropylphenyl)-N,Ndimethylurea); (108), 2,4,6-trinitrotoluene (TNT) C7H5N3O6; (109), tetryl C7H5N5O8 (2,4,6trinitrophenyl-N-methylnitramine); (110) picric acid C6H3N3O7 (2,4,6-trinitrophenol); (111), musk xylene C12H15N3O6 (1-tert-butyl-3,5-dimethyl-2,4,6-trinitrobenzene); (112), nitrofen C12H7Cl2NO3 (1-(4-nitrophenoxy)-2,4-dichlorobenzene); (113), nitroglycerin C3H5N3O9; (114), isosorbide mononitrate C6H9NO6; (115), atrazine C8H14ClN5 (6-chloro-N-ethyl-N0 -isopropyl-1,3,5-triazine2,4-diamine).

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Bio- and Geomacromolecules

Despite occasional claims of an abiotic origin of hydrocarbons occurring in the geosphere, there is no serious controversy that fossil fuels/petroleum are derived from the biomass of formerly living organisms. It is commonly accepted that the various carbon pools in biota exhibit different susceptibility to diagenetic and catagenetic transformation in sedimentary environments. Biogenic organic matter is transformed into geomacromolecules through specific reaction sequences and these geomacromolecules generate petroleum hydrocarbons at elevated temperatures due to deep burial in sedimentary basins. This section provides a brief introduction to bio- and geomacromolecules. Organic substances can be divided roughly into two groups based on their molecular size contrasting low molecular weight compounds from macromolecular substances. From a structural point of view, the description of high molecular weight compounds requires expanded systematics when compared to the low molecular weight compounds, which are sufficiently characterized by their carbon skeleton and the functional groups attached to them. Macromolecular substances exhibit more superordinated structural elements. Besides the chemical characteristics of the monomeric units (the basic building blocks of polymers), the types of recurring linkages that build up the macromolecular skeleton, the degree of polymerization, the extent of cross-linking, and the three dimensional arrangement or orientation of the macromolecular structure, are important parameters for the structural description of macromolecules. Macromolecules represent an essential group of natural compounds with respect to the quantitative distribution on earth, particularly, in living organisms. Their relative contributions to selected biota are summarized in > Fig. 13. Macromolecules with a high degree of order are the physiologically important peptides and proteins, nucleic acids and carbohydrates. Peptides and proteins are formed by multiple linkages of proteinogenic amino acids via amide bonds (so-called peptide bonds), where the number of amino acid moieties distinguishes the relatively small peptides (up to approx. 100 monomers) from proteins (more than approx. 100 monomers). Since amino acids are commonly multifunctionalized molecules they promote the stabilization of superordinated structural assemblies by intramolecular interactions. Hence, the complex structure of proteins is subdivided into primary to quaternary structural elements. The order of amino acids in the protein chain represents the primary structure, which is also called amino acid sequence. Peptide bonds are not symmetrical and, therefore, the direction of the chains is determined by the end members and their free functional groups (the N– and C–terminal groups). Two different forms of secondary structures, the alpha helix and the beta sheet, are dominant in proteins and are fixed by hydrogen bonds. Note that the physico-chemical properties of these structural forms differ significantly and influence the functions of proteins in the organism. The more stable beta sheet structure is realized frequently in proteins with mechanical functions like actin, whereas the more flexible alpha helix is the dominant secondary structure in proteins involved in metabolic and regulatory processes such as enzymes. Note that further types of secondary structures also exist like the triple helix found in collagen. The superordinated folding of protein chains represents the tertiary structure which is stabilized by intramolecular forces like disulfide bridges or weaker interactions. The tertiary structure defines the overall shape of one protein molecule, whereas the quaternary structure refers to the shape of closely associated protein complexes, in which several individual proteins are incorporated.

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. Figure 13 Composition (dry wt%) of main organic constituents (proteins, carbohydrates, lipids and lignin) from various living organisms. Reprinted with permission from Huc (1980).

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The backbone of RNA and DNA molecules is realized by alternating phosphate and ribose or deoxyribose moieties linked via ester bonds. Note that the phosphoric acid is attached to the hydroxyl groups of the adjacent carbohydrate molecules exactly at the 3rd and 5th position. This asymmetric linkage generates a direction of the resulting strains, defined by the position of the terminal hydroxyl groups at the two ends of the strain referred to as 30 or 50 end. Since ribose and deoxyribose are multifunctionalized, they are able to bind another moiety at the 1st position, the heterocyclic nitrogen-containing bases which are of either purine or pyrimidine type. In summary, the individual monomer consisting of one phosphate residue, one sugar molecule, and one heterocyclic base represents a nucleotide, whereas the sub-moiety of carbohydrate and base is called nucleoside. Structural differences between DNA and RNA are due to different carbohydrate units (deoxyribose in DNA or ribose in RNA) and the bases involved (adenine, guanine, cytosine in both, thymine solely in DNA, and uracil solely in RNA). The order of the nucleotides and, in particular, the attached bases represents the sequence, which encodes the genetic information. Like proteins, the individual strains of DNA also build up a secondary structure, the double helix, in which the directions of both the strains are opposite to each other. These helices are stabilized by the so-called base pair interaction, intermolecular forces dominated by hydrogen bonds and related interactions between bases of both strains. Due to its specific shape and connectivity the formation of such base pairs is restricted to certain combinations called complementary base pairs (guanine – cytosine and adenine – thymine/uracil). Note that RNA exists dominantly as mono strains and exhibits a higher variety of secondary structures. A third group of biological macromolecules are the polysaccharides built up by multiple linkages of monomeric sugar moieties (for overview see > Table 6). These monomers represent intramolecular hemiacetals, which are able to stabilize by forming intermolecular acetals through a second addition or elimination reaction with hydroxyl groups of other sugar molecules via so-called glycosidic linkages. Unlike proteins or nucleic acids, most of the polysaccharides are formed solely by one type of monomer resulting in uniform sequences. Most prominent polysaccharides are built up from b-glucose, for example, cellulose or starch. Due to their uniformity in sequence, the differences exist in the network of linkages demonstrating again the interrelationship between molecular structure and physico-chemical as well as the chemical properties. Key aspects of the structural diversity in polysaccharides are the position of hydroxyl groups involved in glycosidic linkages, and the degree of cross-linking. From a structural point of view, the so-called cyclodextrins are an interesting subgroup of oligosaccharides which represent cyclic compounds composed of 7–9 sugar monomers forming a toroid structure, in which lipophilic compounds can easily be captured by inclusion. However, cyclodextrins are nonnatural substances with widespread application in the technosphere. All biological macromolecules described so far are characterized by a high degree of regularity. Further on, the linkages forming the polymeric structures are amide, ester, or acetal structures connecting the monomers via C–O or C–N bonds. The formation of these linkages exhibits a high potential of reversibility, which is an important property with respect to their physiological functions and the required chemical flexibility. However, as a consequence these biological macromolecules also exhibit a low stability outside the living organisms leading to a fast environmental degradation, basically by microbial transformation. An overview of important macromolecules in biota and their diagenetic stability is given in > Table 7. In general, biological macromolecules with lower physiological specialization or relevance exhibit higher diagenetic stability or vice versa. Elevated persistence is usually accompanied by

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. Table 6 Composition of various polysaccharides with respect to their monomers and their glycosidic linkage(s) Biopolymer

Monomer

Position of glycosidic linkage

Cellulose

b-D-Glucose

1–4

Amylose

a-D-Glucose

1–4

Amylopectin

a-D-Glucose

1–4 and 1–6

Glycogen

a-D-Glucose

1–4 and 1–6

Dextran

a-D-Glucan

1–6, 1–3, 1–4, 1–2

Chitin

b-D-Aminoglucose

1–4

Inulin

b-D-Fructose

2–1

Xylan

b-D-Xylose

1–4

. Table 7 Diagenetic stability of certain biomacromolecules according to de Leeuw and Largeau (1993) Biomacromolecules

Occurrence

‘‘Preservation potential’’

Starch

Vascular plants; some algae; bacteria



Glycogen

Animals

 

Cellulose

Vascular plants; some fungi

Pectins

Vascular plants

Chitin

Arthropods; copepods; crustaceans; fungi; algae

+

DNA, RNA

All organisms



Proteins, peptides

All organisms



Cutins, suberins

Vascular plants

+/++

Lignins

Vascular plants

++++

Sporopollenins

Vascular plants

+++

Algaenans

Algae

/+

++++

a lower regularity or order within the macromolecular structures. Interesting examples include suberin and cutin, the two macromolecular substances built up by hydroxylated long chain mono- and dicarboxylic acids via ester bonds. As a result of the multiple functional groups, which can react within each monomer, a high degree of cross-linking is realized in both types of high molecular weight substances. However, the primary structure is irregular and only an average contribution of monomeric units (e.g., o-hydroxypalmitic acid, hexadecanedicarboxylic acid, dihydroxy stearic acids) can be assessed and not the specific regularity of the order of monomers. It is known that besides aliphatic moieties, aromatic structures are incorporated as cinnamic acid derivatives. Besides their different physiological function in plant tissue, both types of macromolecular substances exhibit slight variations in their composition. Cutins exhibit a higher proportion of aliphatic moieties with a slightly lower degree of cross-linking, whereas on the contrary, suberin is characterized by a higher

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aromaticity and extended interlacing. However, knowledge on the three dimensional extension and structure of cutins and suberins is restricted. Note that the biological macromolecules with similar structural properties when compared to suberin and cutin are algaenan and sporopollenin, which are important constituents of algae or spores and pollen, respectively. A further biomacromolecule with a high preservation potential is lignin, the dominant constituent of woody material. The network of lignin molecules is composed irregularly of three building blocks with aromatic and olefinic moieties (p-coumaryl alcohol, coniferyl alcohol, and sinapyl alcohol), which are linked and cross-linked by polymerization at the double bonds or by condensation or substitution at the hydroxyl and ether groups. Crosslinking is also found partially with further biological macromolecules in plants like hemicellulose. Note that the macromolecular backbone is built predominantly via C–C–bonds, and, therefore, is different when compared to the principal mode of linkage in all biomacromolecules discussed so far. The biological function of lignin is associated primarily with the water conduction system in vascular plant stems. Higher stability and also higher lacking of defined structural elements is the characteristic of natural organic macromolecules derived from diagenetic processes. This fraction of organic matter is present in soils and sediments as humic substances which upon further diagenetic and catagenetic processing are transformed into a more decomposed, dispersed, and dominantly amorphous organic material, the so-called kerogen, the latter representing the main form of organic matter in sedimentary rocks. From a structural point of view, lignin and related biomacromolecules with a low rate of decomposition in soils or sediments represent the dominant fraction in humic substances. The molecular structure of humic substances has not been clearly defined, and the common differentiation into the subclasses humins, humic acids, and fulvic acids is based on a procedural definition according to the solubility in alkaline and acidic aqueous solutions. A more detailed structural description of humic substances is restricted to the bulk composition, especially of the dominant functional groups present in these macromolecules (carboxylic acids, hydroxyl and ether groups, quinoid moieties etc.). Additionally, a high proportion of aromatic moieties is evident in many cases. Some structural trends can be associated with the subclasses of humic substances. Following the order from fulvic acids over humic acids to humins, decreasing contents of functional groups and, in particular, oxygen is observed, whereas the carbon content, the degree of cross-linking and the average molecular weight increase. Note that the chemical quality and, consequently, the structural diversity of humic substances depend distinctly on the environment they derive from. Principally, aquatic and terrestrial humic substances and also humic substances from coaly material and soils can be differentiated with respect to their principal structural constitution. Kerogen, the ultimate macromolecular precursor of low-molecular weight petroleum hydrocarbons, represents the major pool of organic matter on earth, but its structural characterization is still limited. Therefore, the common definition of kerogen as a dispersed sedimentary organic material insoluble in organic solvents uses an operational specification but is not based on any structural aspect. The structural diversity of kerogen is enormous but it is obvious, that the major part of kerogen consists of macromolecular compounds with (1) varying contributions of aromatic and aliphatic moieties, (2) various degrees of crosslinking, and (3) diversified functional groups embedded. Its high structural diversity results basically from the multitude of biological precursor substances and the high variability of

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diagenetic modifications. Interestingly, the knowledge on kerogen structures is closely related to the development of analytical techniques. An excellent summary of the present state of knowledge on kerogen structure and properties has recently been published (Vandenbroucke and Largeau, 2007).

8

Physical Properties of Hydrocarbons and Other Organic Compounds

The environmental behavior of an organic compound, like its bioavailability or its tendency to accumulate in certain compartments, is controlled by its macroscopic physico-chemical properties, such as melting and boiling point, density, viscosity, vapor pressure, or aqueous solubility. These properties show systematic variations directly related to the molecular structure of a compound which determines the ability of its molecules to interact with other molecules (or other components of their direct environment such as the surface of solids). Relevant structural features are the presence or absence (as in hydrocarbons) of certain functional groups, intramolecular interactions of different substituents (e.g., shielding effects), or molecular size and shape. The most important intermolecular forces in the order of increasing strength are van der Waals interactions, dipole-dipole interactions, and hydrogen bonding. Van der Waals interactions depend on weak forces between molecules and/or ions, whose energy decreases by the sixth power of the distance between the involved species. Asymmetric distribution of electron density or electronic charge in molecular or ionic species generates permanent dipole moments. A comparison of dipole moments of selected hydrocarbons and functionalized organic compounds illustrates the influence of heteroatoms and functional groups (> Table 8). Dipole moments are typically weak in hydrocarbons. Due to the high degree of structural symmetry certain hydrocarbons such as methane, 2,2-dimethylpropane or benzene do not even possess a permanent dipole moment. Notably, carbonyl compounds (aldehydes and ketones) posses rather elevated dipole moments which contribute to their relatively high melting and boiling points and aqueous solubilities. Hydrogen bonding requires the presence of functional groups to which hydrogen is covalently bound to a heteroatom such as nitrogen or oxygen. Carboxylic acids, alcohols, phenols, or amines are typical compound classes which exhibit the ability to form hydrogen bonds. A negative partial charge is at the more electronegative heteroatom, whereas carbon and hydrogen atoms carry a positive partial charge, in these structural moieties. The resulting dipole moments induce electrostatic forces between functional groups of two different molecules leading to intermolecular interactions, or of the same molecule, resulting in intramolecular interactions. Importantly, functional groups, without hydrogen being bound to a heteroatom such as carbonyl groups, may be involved in hydrogen bonding by interacting with heteroatom-bound hydrogen atoms of functional groups in the same or another molecule. The bond strength ranges from Fig. 14). The difference in boiling points of the most (methane) and

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. Table 8 Dipole moments of selected organic compounds Aliphatic compounds Formula

Name

Aromatic compounds m [D]

CH(CH3)3

Isobutane

0.13

CH3–CH2–CH(CH3)2

Isopentane

C5H8

Cyclopentene

CH2=CH–CH3

Propene

CH2=CH–CH2–CH3 CH2=CH–CH2–CH2– CH3 CH3–CH2–O–CH3

Ethyl methyl ether 1.17

Formula

Name

m [D]

C6H5–CH3

Toluene

0.38

0.13

C6H5–CH2–CH3

Ethylbenzene

0.59

0.20

o-C6H4(CH3)2

o-Xylene

0.64

0.37

C6H5–CH(CH3)2

Isopropylbenzene 0.79

1-Butene

0.44

C6H5–C(CH3)3

tert-Butylbenzene 0.83

1-Pentene

0.50

C12H10

Acenaphthene

0.85

C6H5–O–CH3

Anisole

1.38 1.13

CH3–CH2–NH2

Ethylamine

1.22

C6H5–NH2

Aniline

CH3–CH2–SH

Ethanethiol

1.61

C6H5–SH

Benzenethiol

1.23

CH3–CH2–OH

Ethanol

1.69

C6H5–OH

Phenol

1.22

CH3–(C=O)–O–CH3

Methyl acetate

1.72

C6H5–(C=O)–O– CH3

Methyl benzoate

1.94

CH3–CH2–Cl

Chloroethane

2.05

C6H5–Cl

Chlorobenzene

1.69

CH3–CH2–CHO

Propanal

2.72

C6H5–CHO

Benzaldehyde

3.00

CH3–CH2–(C=O)–CH3

2-Butanone

2.78

C6H5–(C=O)–CH3

Acetophenone

3.02

CH3–CH2–NO2

Nitroethane

3.23

C6H5–NO2

Nitrobenzene

4.22

CH3–CH2–CN

Propionitrile

4.05

C6H5–CN

Benzonitrile

4.18

least (formic acid) volatile C1 compound is as high as 262.5
C. The total range of boiling points for representatives of different compound classes at a given carbon number decreases with increasing carbon number and is below 100
C at C16 for the compound classes depicted in > Fig. 14. Hydrocarbons generally show the lowest, and compound classes such as carboxylic acids and 1-alkanols which are able to form strong hydrogen bonds the highest melting and boiling points. According to Carnelley’s rule high molecular symmetry is associated with high melting points, for example, the melting point of benzene (5.5
C) is significantly higher than that of toluene (94.9
C) despite of the lower molecular weight (Brown and Brown, 2000). Differences in density between members of different compound classes are large for the lower homologues and become systematically smaller with increasing molecular size (> Fig. 15a). As for the boiling points, the largest overall difference among the compound classes depicted in > Fig. 15a is observed between methane and formic acid. The plot also reveals that the density increases with increasing carbon number for hydrocarbons while it decreases for n-alkanoic acids and their methyl esters. Some compound classes such as 1-alkanols, alkanals, and 2-alkanones do not show significant variations of density in relation to the carbon number. At a given carbon number, the densities of n-alkanes and 1-alkenes are relatively similar while those of alkylbenzenes and alkylcyclohexanes are significantly higher. An increasing degree of halogenation of organic compounds leads to a significant increase in density as illustrated for haloethanes in > Fig. 15b; the effect increases in the order

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

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. Figure 14 Boiling points at atmospheric pressure versus carbon number for selected compound classes. All data are from Lide (2002).

fluorine < chlorine < bromine < iodine. Density should not be confused with specific gravity which is calculated as the ratio of the density of a given compound to that of water. In the petroleum industry, the American Petroleum Institute gravity (API gravity) is the most important measure for the quality of crude oils. It is calculated according to the formula API gravity ¼ ð141:5=specific gravity at 15:5
CÞ  131:5; thus API gravity is inversely proportional to density. In-reservoir biodegradation is one of the most important processes leading to a decrease of API gravity and thus a quality deterioration of crude oils. The increase of density in such oils is due to the specific loss of hydrocarbons of relatively low density and a resulting relative enrichment of functionalized organic compounds of higher density. Viscosity is an important physico-chemical property which characterizes the resistance of a fluid being deformed and may be applied to pure organic compounds or mixtures. Within homologous series of compounds it increases exponentially with increasing carbon number as illustrated for selected n-alkanes in > Fig. 16. As a consequence crude oils containing high proportions of long-chain n-alkanes may be highly viscous and therefore difficult to produce. Viscosity decreases with increasing temperature (> Fig. 16); therefore, some crude oils which

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. Figure 15 (a) Density versus carbon number for selected compound classes. (b) Density versus number of halogen atoms for ethane and its halogenated derivatives (for di-, tri- and tetrahaloethanes, data for both isomers were included if available). If available, density data determined at 20
C were used; otherwise data obtained at different temperatures were included as well (total range 162
C to 100
C). All data are from Lide (2002).

are produced as a liquid from their deep hot subsurface reservoirs become solid under surface conditions. For crude oils, a rough positive correlation of API gravity and viscosity is observed. The interaction with different phases in multiphase systems is crucial for the environmental behavior of organic compounds. In natural environments various gaseous (air, natural gas, gas condensates), liquid (aqueous phases, nonaqueous phase liquids NAPL), and solid (inorganic sediment particles and rock matrices, organic particles and amorphous kerogen) phases have to be considered. Very important with respect to the effects of organic compounds on biota is their transport behavior in mobile gaseous and liquid phases. Boiling point and vapor pressure are the relevant properties controlling the concentration of hydrocarbons and other organic compounds in air while aqueous solubility represents a key parameter with respect to their occurrence and distribution in the hydrosphere. Only certain organic compounds are miscible with water. Typically, these consist of rather polar molecules containing functional groups that may form hydrogen bonds. For a broad range of organic compound classes including all known types of hydrocarbons aqueous solubility shows a systematic relationship to specific structural features. In homologous

Hydrocarbons: An Introduction to Structure, Physico-Chemical Properties and Natural Occurrence

1

. Figure 16 Viscosity at different temperatures versus carbon number for selected n-alkanes. All data are from Lide (2002).

series of various compound classes the aqueous solubility decreases with increasing carbon number (> Fig. 17a). It is evident from > Fig. 17a, that the aqueous solubility of hydrocarbons is lower than that of functionalized organic compounds with the same number of carbon atoms. Among the hydrocarbons, acyclic saturated compounds are 1–3 orders of magnitude less water-soluble than aromatic hydrocarbons of similar molecular weight (> Fig. 17b). As a consequence, compounds structurally as different as n-dodecane (C12H26, molecular weight 170) and picene (C22H14, molecular weight 278) have very similar aqueous solubilities at 25
C (n-dodecane 0.00000037 mass %, picene 0.00000025 mass %). The aqueous solubility of cycloalkanes, alkenes, and cycloalkenes is typically slightly higher than that of saturated hydrocarbons of similar molecular weight reflecting their slightly higher polarity. Importantly, some of the mentioned physico-chemical properties show a significant dependence on environmental parameters which has to be taken into account when evaluating the behavior of a specific compound. For example, boiling points show strong pressure dependence while viscosities strongly depend on temperature (> Fig. 16). The phase behavior of hydrocarbon fluids (exclusive occurrence as gas or liquid or co-occurrence of both phases) in subsurface petroleum systems is governed by the given temperature pressure regime. The aqueous solubility of hydrocarbons typically increases with increasing temperature but decreases with increasing salinity. An environmentally highly relevant parameter of organic compounds is their octanolwater partition coefficient, which characterizes the distribution of a compound between 1-octanol (as a model phase for lipophilic compartments) and water phases which are in

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. Figure 17 (a) Aqueous solubility versus carbon number for different compound classes. Formic, acetic, propionic and butanoic acid as well as methanol, ethanol and 1-propanol are miscible with water and therefore do not occur in this plot. (b) Aqueous solubility versus molecular weight for different types of hydrocarbons. With very few exceptions solubility data determined at 25
C were used. All data are from Lide (2002). Please note the logarithmic scale of the y-axes.

equilibrium. Increasing octanol-water partition coefficients as typically seen in homologous series of organic compounds (> Fig. 18) indicate the increasing lipophilic nature of these compounds. At a given carbon number, hydrocarbons and halogenated organic compounds typically show higher octanol-water partition coefficients than (other) functionalized compounds. Octanol-water partition coefficients should be used cautiously when assessing the behavior of a compound in a natural environment. For example, crude oil-water partition coefficients determined for alkyl phenols (Taylor et al., 1997) differ significantly from the octanol-water partition coefficients of these compounds with respect to the differences between isomers. Analogously, Henry’s Law constant may be used to characterize the behavior of gases. According to Henry’s Law, the amount of a given gas dissolved in a given type and volume of liquid is directly proportional to the partial pressure of that gas in equilibrium with that liquid. All physico-chemical data used in this article were taken from the Handbook of Chemistry and Physics (Lide, 2002) which represents a very useful source of information. In Appendix B it also provides a comprehensive compilation of sources of physical and chemical data including web-based resources. It should be noted that in addition to experimental determination of such data modern tools of molecular modeling provide a complementary approach towards an improved understanding of the environmental behavior of hydrocarbon and other organic compounds.

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. Figure 18 Octanol-water partition coefficients versus carbon number for selected compound classes. Please note that log P is used in this plot. All data are from Lide (2002).

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Fiedler H (1996) Sources of PCDD/PCDF and impact on the environment. Chemosphere 32: 55–64. Fleming FF (1999) Nitrile-containing natural products. Nat Prod Reports 16: 597–606. Francke W, Schulz S (1998) pheromones. In: Comprehensive Natural Products Chemistry, vol 8. K Mori (ed.). Oxford: Pergamon, pp. 197–261. Gribble GW (1994) The natural production of chlorinated compounds. Environ Sci Technol 28: 310A–319A. Gribble GW (2000) The natural production of organobromine compounds. Environ Sci Pollut Res Int 7: 37–49. Grice K, Alexander R, Kagi RI (2000) Diamondoid hydrocarbon ratios as indicators of biodegradation in Australian crude oils. Org Geochem 31: 67–73. Hall C, Tharakan P, Hallock J, Cleveland C, Jefferson M (2003) Hydrocarbons and the evolution of human culture. Nature 426: 318–322. Hebting Y, Schaeffer P, Behrens A, Adam P, Schmitt G, Schneckenburger P, Bernasconi SM, Albrecht P (2006) Biomarker evidence for a major preservation pathway of sedimentary organic carbon. Science 312: 1627–1631. Heider J (2007) Adding handles to unhandy substrates: anaerobic hydrocarbon activation mechanisms. Curr Opin Chem Biol 11: 188–194. Hinrichs K-U, Hayes JM, Bach W, Spivack AJ, Hmelo LR, Holm NG, Johnson CG, Sylva SP (2006) Biological formation of ethane and propane in the deep marine subsurface. Proc Natl Acad Sci USA 103: 14684–14689. Huc AY (1980) Origin and formation of organic matter in recent sediments and its relation to kerogen. In: Kerogen, Insoluble Organic Matter from Sedimentary Rocks. B Durand (ed.). Paris: Editions Technip, pp. 445–474. de Leeuw JW, Largeau C (1993) A review of macromolecular organic compounds that comprise living organisms and their role in kerogen, coal and petroleum formation. In: Organic Geochemistry – Principles and Applications. MH Engel, SA Macko (eds.). New York: Plenum Press, pp. 23–72. Lide DR (2002) CRC Handbook of Chemistry and Physics, 83rd edn. 2002–2003. Boca Raton, FL: CRC Press. Mango FD (2000) The origin of light hydrocarbons. Geochim Cosmochim Acta 64: 1265–1277. Marynowski L, Czechowski F, Simoneit BRT (2001) Phenylnaphthalenes and polyphenyls in Palaeozoic

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2 Methods of Hydrocarbon Analysis H. Wilkes Helmholtz Centre Potsdam, GFZ German Research Centre for Geosciences, Organic Geochemistry, Potsdam, Germany [email protected] 1

Introduction – Analytical Challenges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50

2 Extraction, Fractionation, and Separation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 2.1 Liquid Chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 2.2 Gas Chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 3

Identification and Structure Elucidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57

4

Isotope Analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60

5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_2, # Springer-Verlag Berlin Heidelberg, 2010

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Methods of Hydrocarbon Analysis

Abstract: Understanding the interactions of microorganisms and hydrocarbons requires efficient tools for the analysis of the latter in a broad variety of sample matrices. This article provides an overview of key methods used in hydrocarbon analysis. Analytical challenges are related to the structural variability and compositional complexity of naturally occurring hydrocarbons and related heteroatomic compounds. With this in mind important approaches for the extraction, fractionation and separation as well as for the identification and structure elucidation will be discussed. The main emphasis is on liquid and gas chromatographic and on mass spectrometric techniques which are the most important analytical tools in this field of research. Methods of isotope analysis are also covered.

1

Introduction – Analytical Challenges

Our understanding of the behavior and environmental fate of hydrocarbons depends widely on their characterization using state-of-the-art methods of analytical organic chemistry. Key objectives of hydrocarbon analysis are the unequivocal identification and the valid quantitation of individual compounds in a diverse range of matrices. Hydrocarbon analysis plays a particular role in the environmental sciences and in petroleum exploration, production and reformation. Various factors determine the requirements for methods to be used in hydrocarbon analysis. This has to take into account that hydrocarbons occur naturally in great structural diversity and with a significant variability of their physico-chemical properties (for overview See > Chapter 1, Vol. 1, Part 1). They occur and must be determined in nearly all natural matrices including biological tissues, air (e.g., > Chapter 18, Vol. 1, Part 4; Po¨schl, 2005), natural waters (e.g., > Chapter 20, Vol. 1, Part 4), soils (e.g., > Chapter 19, Vol. 1, Part 4) and sediments and sedimentary rocks (including petroleum source and reservoir rocks; e.g., Peters et al., 2005). Typically, hydrocarbons occur in these matrices as complex mixtures, which, moreover, are accompanied by an often even more complex assemblage of heteroatomic compounds. Therefore, analytical procedures are required which allow an efficient separation of hydrocarbons from various types of inorganic components as well as from non-hydrocarbon constituents of natural organic matter. Concentrations of individual constituents in such mixtures may differ by many orders of magnitude. This implies that analytical methods must have an extremely high separation efficiency, allow the unequivocal identification of individual compounds in mixtures often containing numerous other structurally very similar compounds and provide an optimal linearity for the quantitation of individual constituents. Interpretation of analytical data has to consider biases caused by the ‘‘analytical window’’ of the used method. This may be illustrated by the fact that the role of high molecular weight hydrocarbons (>C40) in fossil fuels was only recognized with the advance of high-temperature gas chromatography (e.g., Philp et al., 2004). Only recently, ultrahigh-resolution Fourier transform ion cyclotron resonance mass spectrometry has revealed that crude oil contains organic nitrogen, sulfur and oxygen (NSO) compounds having more than 20,000 distinct elemental compositions (CcHhNnOoSs) (Marshall and Rodgers, 2004). Most of these distinct elemental compositions represent numerous distinguishable, but still unresolved structural isomers. Analytical techniques useful in hydrocarbon analysis evolve rapidly. This contribution will give an introduction to the most important state-of-the-art approaches and highlight some recent developments. It is not within the scope of this article to provide specific technical details of individual analytical methods. This includes all aspects of quantitation such as

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calibration, external versus internal standardization or the consideration of specific response factors of certain compounds or compound classes. If possible, the reader will be referred to recent valuable review articles on the discussed methods. A book on analytical advances for hydrocarbon research has been edited recently (Hsu, 2003). Numerous protocols for the analysis of hydrocarbons have been published by ASTM International, originally known as the American Society for Testing and Materials (http://www.astm.org/index.shtml). The Total Petroleum Hydrocarbon (TPH) working group of the Association for Environmental Health and Sciences has published a useful overview of the analysis of petroleum hydrocarbons in environmental media (http://www.aehs.com/publications/catalog/contents/tph.htm). The Norwegian Industry Guide to Organic Geochemical Analyses (NIGOGA) published by the Norwegian Petroleum Directorate (NPD) provides detailed information on the characterization of geological materials (http://www.npd.no/engelsk/nigoga/default.htm). The NPD also provides the Norwegian Geochemical Standard (NGS) samples (crude oil and source rock samples) as a reference and sets acceptance criteria for selected test parameters. Ample information on analytical methods in general as well as specific techniques used in hydrocarbon analysis can be found in the Encyclopedia of Separation Science. Numerous scientific journals dealing with different aspects of analytical chemistry regularly publish review articles on new technical developments.

2

Extraction, Fractionation, and Separation

It is extremely important that any type of sample to be analyzed for hydrocarbons is collected, handled and stored prior to analysis under appropriate conditions which avoid post-sampling alteration of the composition of indigenous hydrocarbons as well as contamination by hydrocarbons from external sources. Even samples of consolidated sedimentary rocks may be deeply penetrated by hydrocarbons or other contaminants upon exposure to sources of such compounds (Brocks et al., 2008). Any sample handling should avoid direct or indirect contact to materials which might be sources of hydrocarbons, particularly plastics. Important measures to avoid post-sampling alteration include storage in the dark at low temperature under an inert oxygen-free atmosphere and sterilization to eliminate biological activity (particularly in water or water-containing samples). The analysis of volatile organic compounds requires sample storage in gas-tight vials. A multitude of methods is used for the extraction of hydrocarbons from diverse matrices (Bicking, 2000; Smith, 2003). Analytical extraction is based on the two- or multi-phase equilibration of the target compound(s) between the sample matrix and the extractant which is influenced by numerous factors such as temperature, pH, ionic strength, time, repetition, surface area and stirring/mixing (Bicking, 2000). According to the principle ‘‘like dissolves like’’ mainly nonpolar liquid or solid extractants are use in hydrocarbon analysis. This implies that most extraction techniques will co-extract other compound classes of similar polarity. It may be generally differentiated between liquid-liquid and liquid-solid extraction techniques. Liquid-liquid extraction using separatory funnels or continuous liquid-liquid extractors is useful for extraction of hydrocarbons from aqueous samples. Extraction of total lipids including hydrocarbons from biological tissues is accomplished by the method of Bligh and Dyer (1959). Liquid-solid extraction techniques have a broader application range. Soxhlet techniques are commonly used for the extraction of hydrocarbons and other types of organic compounds from solid matrices such as sediment and rock samples. Accelerated

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solvent extraction (ASE) is an automatable technique, which combines elevated temperatures and pressures with liquid solvents (Richter et al., 1996). Supercritical fluids, particularly carbon dioxide can be used as an environmentally friendly solvent for extraction of hydrocarbons, crude oil and related products from solid samples (Rudzinski and Aminabhavi, 2000). Solid-phase micro extraction (SPME) is a convenient tool for rapid sample preparation to analyze hydrocarbons in gaseous, liquid and solid phases (e.g., Tang and Isacsson, 2008). As SPME is a non-exhaustive extraction technique in which only a small portion of the target analyte is removed from the sample matrix, calibration for quantitative analysis is very important (Ouyang and Pawliszyn, 2008).

2.1

Liquid Chromatography

Soluble organic matter (SOM = crude oil, non-aqueous phase liquids or solvent extracts obtained by the above mentioned methods using liquid extractants) typically represents a highly complex mixture of numerous compound classes which in most cases requires specific clean-up procedures prior to instrumental analysis. Mainly liquid chromatography (LC) is used for the group-type analysis of SOM or to provide chemically defined sub-fractions suitable for further characterization e.g., by gas chromatography. An excellent overview of liquid chromatographic methods used in the separation of petroleum and related products has been published by Barman et al. (2000). In most cases solvent extracts have to be concentrated prior to analytical or preparative LC; caution is required during this step to avoid the loss of volatile organic compounds. Hydrocarbon analysis employs various techniques of liquid chromatography such as thinlayer chromatography (TLC), open-column liquid chromatography, medium pressure liquid chromatography (MPLC), high-performance liquid chromatography (HPLC) and supercritical fluid chromatography (SFC). Both normal- and reversed-phase applications are used, however, due to the nonpolar nature of the analytes normal-phase LC is more common in hydrocarbon analysis than in other fields of organic-chemical analysis. Normal phases such as silica or alumina are characterized by polar surfaces while reversed phases typically are modified silica gels which due to the presence of chemically-bonded hydrophobic groups are characterized by a nonpolar surface. Accordingly, solvents or solvent systems used for isocratic or gradient elution are typically nonpolar in normal- and polar in reversed-phase separations; commonly used solvents are listed according to increasing eluotropic strength in a so-called eluotropic series. SOM is easily separated into compound classes by LC methods. This includes the separation of hydrocarbons from heteroatom-containing compounds as well as the further separation of the hydrocarbon fractions into more specific sub-classes. The so-called asphaltenes are a sub-fraction of the non-hydrocarbons in SOM which is insoluble in nonpolar solvents; accordingly, asphaltene precipitation is a standard procedure in sample preparation prior to LC separation (e.g., Theuerkorn et al., 2008). Automatable routine methods such as MPLC provide a rapid and convenient separation of the so-called maltenes, i.e., the asphaltene-free fraction of SOM, into saturated hydrocarbons, aromatic hydrocarbons and NSO compounds (= resins) (e.g., Radke et al., 1980). Specific methods may be used for further fractionation of the resins into chemically well-defined sub-fractions (e.g., Willsch et al., 1997). Both the saturated and the aromatic hydrocarbon fractions obtained by such procedures are directly usable for further analysis by gas chromatography (see below). However, both fractions still consist of highly complex mixtures of structurally diverse constituents.

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The so-called saturated fractions are principally composed of three structural types, the nalkanes, the branched alkanes and the cycloalkanes. Furthermore, depending on the presence of such compounds in a given sample and on the analytical conditions, the saturate fraction may contain unsaturated hydrocarbons (alkenes) with one or even more double bonds. In many cases, n-alkanes represent the main constituents; however, depending on the sample origin and history other compound types may be predominant, for example as a result of biodegradation processes in natural environments. Certain types of polycyclic hydrocarbons derived from biogenic precursors such as steroids or triterpenoids, known as the so-called biomarkers, typically occur in trace amounts in fossil organic matter but have a high information content (for details see Peters et al., 2005). The separation of these compound classes is challenging due to the small differences in physico-chemical properties and requires specific methods such as the use of zeolite molecular sieves (Peters et al., 2005). Likewise, the so-called aromatic hydrocarbon fractions are highly complex mixtures of compound types. The aromatic hydrocarbons sensu stricto may be classified according to the number of annulated aromatic rings as well as to the degree of alkylation. Again, depending on the presence of such compounds and the analytical conditions, unsaturated hydrocarbons may be present. Furthermore, aromatic hydrocarbon fractions may contain certain types of rather nonpolar non-hydrocarbons such as thiophenes, benzothiophenes, dibenzothiophenes and the oxygen-analogous aromatic heterocycles (See > Chapter 1, Vol. 1, Part 1), which due to similar physico-chemical properties are difficult to separate from aromatic hydrocarbons and often are analyzed together using compound-selective detection methods such as gas chromatography-mass spectrometry (see below). The sulfur aromatic heterocycles may be separated from aromatic hydrocarbons by ligand exchange chromatography (LEC) using PdCl2 and CuCl2 on silica gel columns (see Rudzinski et al., 2005 and reference therein). Various methods have been described to separate aromatic hydrocarbons according to ring number (e.g., Radke et al., 1984). A broad variety of detectors may be used in LC of hydrocarbons, including spectroscopic (UV-Vis, fluorescence, infrared), bulk property (refractive index, evaporative light-scattering, dielectric constant, flame-ionization), mass spectrometric and element specific detectors. It has to be taken into account that different detection principles may have specific advantages and disadvantages. E.g., UV-Vis detection provides excellent sensitivity for aromatic but is not applicable to saturated hydrocarbons. The utility of common atmospheric pressure ionization interfaces used for on-line coupling of LC and mass spectrometry in the analysis of volatile and/or nonpolar compounds is rather limited. TLC coupled to flame-ionization detection is an important compound group screening method in hydrocarbon analysis.

2.2

Gas Chromatography

Most environmentally significant hydrocarbons are relatively volatile and – due to their predominant origin from fossil fuels – thermally rather stable. Therefore, gas chromatography (GC), which requires that the target compounds may be vaporized without destruction, has been established as the most important method for the separation of hydrocarbons on the molecular level. The use of GC in the analysis of hydrocarbons and related compounds, e.g., persistent halogenated organic pollutants has been reviewed for example by Barman et al. (2000), Beens and Brinkmann (2000), Philp (2000), Blomberg et al. (2002), Santos and Galceran (2002) and van Leeuwen and de Boer (2008).

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GC is based on two-phase fractionation of a mixture of compounds between a liquid stationary (typically a high-boiling liquid) and a gaseous mobile (mostly helium) phase. Separation of analyte molecules is controlled by their partition behavior between the two phases and by their vapor pressure. Gas chromatographs consist of three main physical components, an inlet system, an analytical column (typically mounted in a temperatureprogramable oven) and a detector system. GC analysis of gases and highly volatile organic compounds is accomplished using gas source inlets, purge and trap systems or headspace samplers. Liquid samples may be injected via syringes through split/splitless, on-column or programed temperature vaporizing (PTV) injector systems (for a review of sampling systems see Davies, 2000). Relatively new techniques allow the on-line coupling of sorbent- and liquidphase micro extraction from solid and liquid samples with GC analysis for various types of compounds (Hyo¨tyla¨inen, 2008; Hyo¨tyla¨inen and Riekkola, 2008). The most common detector systems used in GC are the flame ionization detector (FID), the thermal conductivity detector (TCD) and different types of mass selective detectors (MSD, see also following section). In addition, various types of element- or compound-specific detectors are used, for example the electron capture detector (ECD) which is highly sensitive towards halogenated organic compounds. Efficient separation of diverse target compounds is enabled by the proper choice of analytical columns and oven temperature programs. As an example gas chromatograms of crude oil samples which are biodegraded to different extents are shown in > Fig. 1. In modern GC typically fused silica capillary columns coated with different types of stationary phases are used. Most common in hydrocarbon analysis are nonpolar stationary phases such as pure polydimethylsiloxanes or slightly more polar polysiloxanes in which a certain proportion (e.g., 5%) of the methyl groups is substituted by phenyl groups. Important physical parameters influencing the separation characteristics of the analytical columns are the column length, its inner diameter and the film thickness of the stationary phase. The observed GC retention times depend on several factors, e.g., temperature, flow rate and column length and thus are not an ideal parameter for identification purposes; instead, retention indices may be useful for identifying compounds in complex mixtures (Song et al., 2003). Optimized oven temperature programs may be required to properly resolve specific target compounds such as diastereomers with very similar physical properties (see > Fig. 2 for an example from the recent literature). Enantioselective GC using chiral stationary phases enables the separation of enantiomers (see Schurig, 2002 for a general review and Bastow et al., 1998 for an application to hydrocarbons in geological samples). It should be mentioned that the gas chromatographic behavior of various classes of non-hydrocarbons, e.g., carboxylic acids, may be significantly improved by chemical derivatization prior to GC analysis (for a review of important derivatization methods see Husek, 2000). Preparative capillary GC is useful for the isolation of individual hydrocarbons even from unresolved complex hydrocarbon mixtures of biodegraded crude oil (Sutton et al., 2005). High-temperature and comprehensive two-dimensional GC can be identified as two relatively recent methodological advances which may contribute significantly to an improved analytical characterization of complex mixtures of hydrocarbons. High-temperature GC is a key technique in extending the molecular application range of gas chromatography (e.g., Kaal and Janssen, 2008) which mainly benefited from the development of better temperature-resistant GC columns. Characterization of crude oils by high-temperature GC has provided new insights into the composition and properties of petroleum with a strong impact on exploration and production (Philp et al., 2004); an example is shown in > Fig. 3.

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. Figure 1 Thermovaporization whole oil gas chromatograms for four crude oil samples from petroleum reservoirs offshore Angola. The extent of biodegradation in these oils increases from top to bottom. This is recognized by the decreasing amounts of resolved compounds such as n-alkanes (numbers indicate carbon chain length) and the increase of an unresolved complex mixture (UCM). The UCM represents a specific analytical challenge in the context of modern hydrocarbon research. API is the most important bulk parameter to characterize crude oil quality; it decreases with increasing extent of biodegradation. The pristane/n-heptadecane (Pr/n-C17) and phytane/ n-octadecane (Ph/n-C18) ratios are commonly used to assess variations in the extent of biodegradation. Reprinted from Elias et al. (2007), with permission from Elsevier.

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. Figure 2 Partial m/z 57 mass chromatogram showing separation of the diastereomers of three acyclic isoprenoidal hydrocarbons in standard diesel fuel. nC17 and nC18 are n-alkanes and IS = internal standard (d10-anthracene). The separation of the diastereomers of norpristane (2,6,10-trimethylpentadecane), pristane (2,6,10,14-tetramethypentadecane) and phytane (2,6,10,14-tetramethyhexadecane) was achieved using an unusual oven temperature program of an initial temperature of 50 C, then ramped at 1 C/min to 145 C and held for 60 min, then ramped at 5 C/min to 320 C. The method was applied to determine the extent of biodegradation of fuels in soils from a bioremediation trial at Casey Station, Antarctica. The biological diastereomers of pristane (meso; RS = SR) were depleted more rapidly during moderate biodegradation than the geological or mature diastereomers (RR and SS). Reprinted with permission from McIntyre et al. (2007). Copyright (2007) American Chemical Society.

. Figure 3 High-temperature gas chromatogram (HTGC) of the solid deposit collected from an oil storage tank. The predominance of the high molecular weight hydrocarbons (HMWHCs) in the C30–C60 region of the chromatogram resulted from the accumulation of precipitated HMWHCs over time in the storage tank. Reprinted from Boukadi et al. (2005) with permission from Elsevier.

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In comprehensive two-dimensional gas chromatography (GC  GC) the entire sample is subjected to two distinct analytical separations resulting in an enhanced separating capacity most useful for the characterization of complex mixtures of organic compounds (for recent reviews see Adahchour et al., 2008; Dallu¨ge et al., 2003; Pierce et al., 2008; Vendeuvre et al., 2007). GC  GC may become even more powerful by on-line coupling to mass spectrometry (e.g., Mondello et al., 2008).

3

Identification and Structure Elucidation

A broad range of methods is used for the structural characterization of organic compounds including but not limited to elemental analysis, spectroscopic methods (infrared (IR), Raman, UV/Vis, nuclear magnetic resonance (NMR) and electron spin resonance (ESR) spectroscopy), X-ray crystallography and mass spectrometry (MS). Although several of these methods can be coupled on-line to LC and/or GC, MS is by far prevailing in the so-called hyphenated techniques commonly used in hydrocarbon analysis. It is adequate to point out that rigid structure elucidation of individual organic compounds requires well-defined criteria to be fulfilled. Most importantly mass spectrometry alone will generally not be sufficient for an unequivocal identification of a specific compound. Even high-resolution mass spectral data confirming the elemental composition of a molecular species do not allow the differentiation of structurally related isomers. Two principal strategies may be pursued in order to gain an unambiguous identification. The first is to isolate the compound in question in pure form and then apply methods such as NMR spectroscopy or X-ray crystallography, which allow an unequivocal structure assignment. The second is to deduce a structure hypothesis from the available analytical data, synthesize an authentic standard and demonstrate that it has the same properties as the target compound (mass spectrum, retention behavior in LC and/or GC etc.). Both strategies have been successfully applied to the structure elucidation of complex hydrocarbons and related heteroatomic compounds in natural matrices (e.g., Achitouv et al., 2004; Behrens et al., 1998; Belt et al., 2000; Dahl et al., 2003; Grosjean et al., 2000; Kenig et al., 2003; Rouquette et al., 2005; Shetty et al., 1994; Sinninghe Damste´ et al., 2005). The particular importance of MS for hydrocarbon analysis is related to the opportunity to detect numerous individual molecular species in a complex mixture in parallel. On-line coupling with modern separation techniques provides an almost unlimited separation power. Mass spectrometers consist of three main physical components, an ion generating system (the ion source), a mass analyzer and a detector system. Different types of time-offlight, beam-type and ion-trapping mass analyzers are used (McLuckey and Wells, 2001). It has to be taken into account that different compound types may require specific methods of ion generation depending on structure, molecular weight and polarity. The physicochemical properties of the target compounds determine whether they may be introduced into the ion source in gaseous, liquid or solid form (e.g., Vestal, 2001). Historically, volatile hydrocarbons have been analyzed predominantly by GC-MS. The GC column is directly fitted into the ion source of the mass spectrometer via a transfer line. This technique uses mainly electron ionization (EI) and to a lesser extent chemical ionization (CI). In EI mode, a beam of electrons is directed into molecular vapor at reduced pressure, one or more secondary electrons are ejected, and relatively unstable odd-electron or multiply charged positive ions are produced (Vestal, 2001). The instability of these ions is due to the high electron energy of typically 70 eV which induces specific fragmentation reactions. As these fragment ions are predominantly

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formed in the ion source, i.e., prior to mass analysis, an EI mass spectrum represents a very useful source of information on the structure of the target compound. Many other common ionization methods generate predominantly molecular or pseudo molecular ions which are either positively or negatively charged. Here, tandem MS technologies are a meaningful approach to define the mass-to-charge relationship between a precursor ion and a product ion (McLuckey and Wells, 2001) and thus to obtain valuable structural information. Tandem MS techniques are for example a key method in analyzing hydrocarbon biomarkers occurring in trace amounts in natural organic matter (Peters et al., 2005). Different modes of mass spectral analysis including full scans to obtain the most comprehensive structural information, selected ion monitoring (SIM) for highly sensitive detection of certain molecular species or multiple reaction monitoring (MRM) to achieve an optimal selectivity may be used both in GC- and LC-MS applications. The mass resolution required for an unambiguous determination of the elemental composition depends on the mass difference between the molecular species in question and increases with increasing molecular weight. Very important progress with respect to the comprehensive compositional characterization of complex mixtures of hydrocarbons and associated non-hydrocarbons comes from the advances in high- and ultrahigh-resolution MS, particularly Fourier transform-ion cyclotron resonance-mass spectrometry (FT-ICR-MS; Marshall and Rodgers, 2004; Sleighter and Hatcher, 2007). Application examples of FT-ICR-MS specifically relevant to hydrocarbon microbiology are the demonstration of complete incorporation of hydrocarbons into membrane phospholipids from Rhodococcus hydrocarbon degrading bacteria (Rodgers et al., 2000) and the assessment of subsurface anaerobic microbial alteration of the acidic and neutral polar NSO compounds in crude oil (Kim et al., 2005; > Fig. 4). An excellent general overview of MS applications in the environmental sciences has been provided by Richardson (2001). GC-MS is one of the most attractive and powerful techniques for routine analysis of organic pollutants including hydrocarbons in environmental media as well as the comprehensive compositional characterization of petroleum and related products due to its good sensitivity and high selectivity and versatility (e.g., Peters et al., 2005; Santos and Galceran, 2003). Atmospheric pressure ionization (API) techniques enable the on-line coupling of other separation methods including HPLC, SFC and electro analytical methods to MS, thus allowing the characterization of organic species that are not amenable to GC analysis (Tomer, 2001). The most common API technique, electrospray ionization (ESI) is well-suited for the analysis of polar organic compounds which readily form ions in solution. More appropriate for the analysis of hydrocarbons sensu stricto – in particular those of higher molecular weight such as large polycyclic aromatic hydrocarbons – are atmospheric pressure chemical ionization (APCI) and, to a lesser extent, atmospheric pressure photo ionization (APPI). The book edited by Hsu (2003) contains several useful articles on the characterization of petroleum and its fractions using MS techniques. Herod et al. (2007) recently reviewed analytical methods that have been developed for characterizing complex liquid mixtures derived from fossil fuels with a special emphasis on advances in the analysis of increasingly heavier materials. Biogeomacromolecules play the key role in the transformation pathway from biomass to kerogen, which is the ultimate precursor of oil and gas formed by geothermal reactions deep within sedimentary basins. The structures of resistant biomacromolecules and geomacromolecules are less regular than those of important but rather labile biopolymers such as carbohydrates, proteins and nucleic acids. Their structural characterization therefore bears a great challenge for analytical chemistry. Our present knowledge on the structure, composition and behavior of such macromolecular organic matter has been the subject of excellent recent

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. Figure 4 Broadband negative-ion electrospray ionization 9.4 T FT-ICR mass spectra of crude oils at distinct stages of biodegradation: non-degraded (0), moderately degraded (2), heavily degraded (4) and very severely degraded (6) genetically related oils. To show the complexity of the spectra, one mass scale-expanded segment is displayed at the top of the diagram. Reprinted from Kim et al. (2005), with permission from Elsevier.

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review articles (De Leeuw et al., 2006; Vandenbroucke and Largeau, 2007). Numerous techniques are being used in the analytical characterization of bio- and geomacromolecules including methods such as IR and NMR spectroscopy, however, thermal analysis appears to be the most important approach for a detailed analysis of these materials on the molecular level. Pyrolysis coupled on-line to GC and GC-MS enables the direct determination of the occurrence of certain building blocks in the macromolecules (e.g., Philp, 2003; Sobeih et al., 2008). Thermally assisted hydrolysis and methylation using specific reagents has a strong potential to provide additional chemical structure information (Challinor, 2001). Beyond structural analysis, pyrolytic methods are an important tool to simulate the formation of oil and gas from kerogen (e.g., Horsfield et al., 1989; Lafargue et al., 1998). Thermal as well as numerous other techniques including chemical degradation methods are also important tools to characterize organic compound bound residues in soil and sediment (Northcott and Jones, 2000).

4

Isotope Analysis

Isotope analysis has been an important tool in biogeochemistry and petroleum geochemistry for more than 50 years (for a recent review see Galimov, 2006). Its use is based on small but significant variations in the natural abundance of stable isotopes of various elements in different materials. Several articles in this Handbook discuss the application of stable isotopes towards understanding the origin and fate of hydrocarbons in natural environments and their interactions with microorganisms (See > Chapter 49, Vol. 5, Part 3; > Chapter 20, Vol. 5, Part 2; > Chapter 5, Vol. 1, Part 2). The principal tool to determine stable isotope ratios of elements of interest in organic matter is isotope ratio mass spectrometry (IRMS). Historically, the determination of the exact isotopic masses was a relatively early achievement while the accurate determination of isotopic abundances remained a problem and advanced significantly only with the introduction of dual collectors and compensation measurements (Budzikiewicz and Grigsby, 2006). Important aspects of modern high precision isotope ratio monitoring techniques in mass spectrometry are discussed in a number of review articles (Brand, 1996; Coplen et al., 2006; Werner and Brand, 2001). The breakthrough in the application of stable isotopes in hydrocarbon-related research came with the introduction of rapid routine methods for compound specific isotope analysis (CSIA). For the determination of carbon isotopes of volatile organic compounds, a GC is coupled to an IRMS via a small-volume combustion furnace, in which the organic carbon is quantitatively converted to carbon dioxide, and a unit allowing removal of water from the gas stream by diffusion through a selective membrane (Freeman et al., 1990; Hayes et al., 1990). Subsequently, the compound specific determination of hydrogen isotope ratios became possible by the introduction of an interface system which enables quantitative conversion of hydrogen bound in organic compounds to molecular hydrogen by pyrolysis at temperatures >1,400 C (Hilkert et al., 1999). A state-of-the-art review on isotope-ratio detection for gas chromatography has been provided by Sessions (2006). An example of the primary data obtained by GC-IRMS for the determination of carbon isotope ratios of n-alkanes is shown in > Fig. 5. More recently, approaches for on-line coupling of LC and IRMS (Krummen et al., 2004) and carbon isotopic analyses of nanogram quantities of nonvolatile organic carbon (Sessions et al., 2005) have been described. Almost two decades of compound specific isotope analysis have revealed the enormous potential of this technique, particularly for the characterization of hydrocarbons. Applications

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. Figure 5 A typical GC-IRMS trace of m/z 44 (a), ratio of m/z 45–44 (b), and d13C (c) values for a homologous series of n-alkanes from a marine aerosol sample collected at Chichi-Jima, a remote island in the western North Pacific on 24–27 April 1990. IS, internal standard (C20 fatty acid methyl ester). The signal of m/z 45 is multiplied by 100 times. Reprinted from Bendle et al. (2006), with permission from Elsevier.

include but are not limited to the investigation of biosynthetic pathways (e.g., Hayes, 1993; Sessions et al., 1999), the monitoring of biodegradation in contaminated aquifers and other polluted environments (e.g., Meckenstock et al., 2004; Philp et al., 2007), the quantification of sequestration of specific organic compounds in soil (e.g., Glaser, 2005), the understanding of origin and degradation processes of petroleum (e.g., > Chapter 5, Vol. 1, Part 2) or the reconstruction of past environments and climates (e.g., Huang et al., 2001; Sachse et al., 2004).

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Beside the determination of stable isotopes, also natural abundance molecular-level radiocarbon analysis using accelerator mass spectrometers appears to have a strong but not yet fully explored potential as a tool to trace the cycling of organic compounds in the environment. Pure organic compounds required for radiocarbon analysis may be isolated from complex mixtures by preparative GC (Eglinton et al., 1997) or HPLC (Smittenberg et al., 2002). Molecular-level radiocarbon analysis has been used for example to demonstrate microbial assimilation of ancient organic carbon during shale weathering (Petsch et al., 2001), to determine microbial carbon sources in petroleum contaminated sediments (Slater et al., 2005) or to characterize the buildup of refractory organic carbon in boreal soils (Smittenberg et al., 2006).

5

Research Needs

Recent methodological advances, such as high temperature gas chromatography, Fourier transform-ion cyclotron resonance-mass spectrometry or compound specific isotope analysis to mention just three, provide a strongly improved separation efficiency and resolution for the analysis of mixtures of organic compounds. It is only now that we begin to realize the true complexity of assemblages of hydrocarbons and related heteroatomic compounds in natural environments. The future must show to which extent this progress in compositional characterization of organic matter can also contribute to an improved understanding of processes in which hydrocarbons and microorganisms interact. Desirable achievements in analytical methods include but are not limited to the overcoming of biases in quantitation which are due to different response of various compound classes (for example those observed in atmospheric pressure ionization techniques), the isomeric differentiation of molecular species identified by high-resolution mass spectrometry or routinely applicable tools to asses the intramolecular isotopic composition of organic compounds.

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Behrens A, Wilkes H, Schaeffer P, Clegg H, Albrecht P (1998) Molecular characterization of organic matter in sediments from the Keg River formation (Elk Point group), western Canada sedimentary basin. Org Geochem 29: 1905–1920. Belt ST, Allard WG, Masse G, Robert J-M, Rowland SJ (2000) Highly branched isoprenoids (HBIs): identification of the most common and abundant sedimentary isomers. Geochim Cosmochim Acta 64: 3839–3851. Bendle JA, Kawamura K, Yamazaki K (2006) Seasonal changes in stable carbon isotopic composition of n-alkanes in the marine aerosols from the Western North Pacific: Implications for the source and atmospheric transport. Geochim Cosmochim Acta 70: 13–26. Bicking MKL (2000) Analytical extractions. In Encyclopedia of Separation Science. ID Wilson (ed.). Oxford: Academic Press, pp. 1371–1382.

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3 Natural Gas Hydrates J. M. Schicks Helmholtz Centre Potsdam – GFZ German Research Centre for Geosciences, Section 4.2, Potsdam, Germany [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68 1.1 Gas Hydrates in Nature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 68 2

Structure and Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69

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Hydrate Formation Processes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 71

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Thermodynamic Properties and Hydrate Formation Kinetics . . . . . . . . . . . . . . . . . . . . 74

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 76

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_3, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: In this chapter the characteristics of clathrate hydrates of natural gases, generally called gas hydrates, will be presented. Starting with an introduction to hydrate structures, which have been verified in nature as well as the associated hydrate formers, the phase diagrams exhibiting the stability fields and thermodynamic properties of these natural systems with respect to their composition will be discussed. Natural gas hydrates are methane-rich but may also contain CO2, H2S and other hydrocarbons and hence vary in their thermodynamic properties. Different models regarding the formation and growth processes, including kinetics with respect to heat and mass transfer effects, experimental observations regarding the cage occupancy during the formation process as well as the influence of sediments and pore water salinity will be presented and discussed.

1

Introduction

1.1

Gas Hydrates in Nature

Gas hydrates are solid, ice-like substances composed of water and gas molecules. They occur in nature at elevated pressures, low temperatures and when sufficient amounts of gas and water are available. These conditions are given at the seafloor and in permafrost regions. Therefore gas hydrate occurrences can be verified worldwide – in oceanic sediments on the continental shelves and slopes along active and passive margins and regions with similar conditions, such as the Black Sea or the Caspian Sea and also in polar sediments and permafrost areas (such as in the Canadian arctic or Siberia) (See > Chapter 12, Vol. 1, Part 3) (Cherskiy et al., 1985; Dallimore et al., 1999; Kvenvolden and Lorenson, 2001). In marine sediments gas hydrates can be found in depth up to 1,100 m depending on the p-T-conditions, while gas hydrates in polar continental regions may be found in depth between 150 and 2,000m (Kvenvolden and Lorenson, 2001). Natural gas hydrates encase predominantly methane, but also higher hydrocarbons as well as CO2 and H2S. The amount of additional gases beside methane varies from less than 1 mol% (e.g., Black Sea) to more than 40 mol% (e.g., Caspian Sea, Gulf of Mexico) (Kvenvolden and Lorenson, 2001). Investigations on hydrocarbon gases in natural gas hydrates with respect to the carbon-isotopic (d13C) composition provide data for the interpretation of the origin of these gases. Analyses of marine gas hydrates show that most of these samples have a carbonisotopic composition of methane lighter than 60‰ (relative to the Peedee Belemnite standard) indicating a microbial methane origin. It is very likely that this microbial methane is a product of CO2 reduction from organic matter to methane as a result of methanogenic processes occurring in shallow sediments (Kvenvolden, 1995; Kvenvolden and Lorenson, 2001). A carbon-isotopic composition above 50‰ and the occurrence of higher hydrocarbons such as ethane or propane in the hydrate indicates a thermogenic gas origin (Bernard et al., 1976; Schoell, 1988). These gases result from thermal decomposition of organic matter in sediment depths generally greater than 1,000 m. H2S, which is occasionally incorporated in marine gas hydrates occurring in shallow sediments above SMI (sulfate-methane-interface) is here locally produced by the reduction of sulfate via anaerobic oxidation of methane (AOM) as a result of a complex interaction of microbes which use the sulfate to oxidize the methane anaerobically (See > Chapter 12, Vol. 1, Part 3) (Barnes and Goldberg, 1976; Boetius et al., 2000; Kastner et al., 1998; Zehnder and Brock, 1979).

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. Figure 1 Nodules of natural gas hydrate embedded in sediment. Sample was recovered during the IODP Expedition 311.

Gas hydrates occur finely disseminated or nodular, in veins or as massive layers in the sediments. > Fig. 1 presents a sample of hydrate bearing sediments which was recovered during the IODP Expedition 311, showing nodules of gas hydrate embedded in the sediment. Disseminated hydrates represent the large majority of marine gas hydrates and may dissociate rapidly if p-T-conditions change (Sloan and Koh, 2008). Depending on the formation conditions, hydrates exhibit an either massive or porous habitus, e.g., as demonstrated in natural gas hydrate samples from the Cascadia Margin: in the presence of free gas, a porous hydrate may grow downward towards rising gas (methane) bubbles (Suess et al., 2001). Huge amounts of gas can be stored in gas hydrate deposits: 1 m3 gas hydrate releases approximately 164 m3 gas at standard p-T-condition. The global estimates of hydrate-bound gas in marine sediments are highly speculative and range between less than 1  1015 m3 to more than 3,000  1015 m3 (Milkov, 2003). These variations in calculated results are caused by the different assumptions regarding the occurrence and composition of natural gas hydrates. Depending on the composition of the feed gas, the formation and growth processes of the hydrates as well as the stability field of the resulting hydrate phase vary. This will be discussed in the following paragraphs.

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Structure and Composition

Gas hydrates are non-stoichiometric, crystalline solids. Water molecules arrange in a threedimensional, hydrogen bonded framework with defined cavities. These cavities are occupied by gas molecules which prevent the water cages from collapsing. From the chemical point of view gas hydrates are assigned to clathrates.

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. Table 1 Number of cavities per unit cell for different gas hydrate structures (Sloan, 1998) Cavities 512

51262

51264

435663

51268

Structure I

2

6

–

–

–

Structure II

16

–

8

–

–

Structure H

3

–‘

–

2

1

Three hydrate structures are known to be capable of holding light hydrocarbons and have been confirmed in natural gas hydrate occurrences: the cubic structures I (sI) and II (sII) and the hexagonal structure H (sH). All structures are composed of various kinds of cages (> Table 1). In structure I two small pentagonal dodecahedrons (512) are combined with six tetrakaidecahedrons (51262) into a unit cell. The pentagonal dodecahedron consists of 20 water molecules forming a 12-sided cavity which has pentagonal faces with equal angles and edge length. It is the smallest cavity type with an average radius of 0.39 nm and part of all hydrate structures. The tetrakaidecahedron combines 12 pentagons with 2 hexagons and, therefore, the radius of these cavities increases to 0.433 nm (McMullan and Jeffrey, 1965; Sloan, 1998) (> Fig 2). A unit cell of structure II consists of 16 pentagonal dodecahedrons (512) and 8 hexakaidecahedrons (51264). The latter cage combines 12 pentagonal faces with 4 hexagonal faces and has a radius of about 0.473 nm (Mak and McMullan, 1965; Sloan, 1998). Structure H is the only structure containing a cavity with three square faces in addition to pentagonal and hexagonal faces (435663). The combination of three pentagonal dodecahedrons (512), two irregular dodecahedrons (435663) and one icosahedron 51268, which is the largest cavity (average radius 0.579 nm), forms the characteristic unit cell of structure H (Ripmeester et al., 1987; Sloan, 1998). The formed structure depends – amongst others – on the composition of the feed gas, namely the size of the enclathrated gas molecules: small guest molecules, such as nitrogen or oxygen form structure II hydrates with both large and small cavities being filled. Slightly larger guest molecules such as methane, CO2 or H2S form structure I hydrates, with partial filling of the small cavities, whereas larger molecules such as propane form structure II hydrates with small cavities being empty. Even larger molecules such as neo-hexane form structure H hydrates in the presence of a supporting gas that fills smaller cavities of the structure (e.g., methane). However, in the presence of a gas mixture, the relationship between structure and size is not always straightforward. Gas molecules which individually form structure I hydrates may form structure II in a mixture, as has been demonstrated for methane-ethane hydrates in certain proportions (Subramanian et al., 2000). All three structures have been confirmed on natural hydrate samples. Recently, oceanic gas hydrates on the Cascadian margin have been investigated for the first time in situ using Raman spectroscopy. They have been identified as structure I hydrates containing predominately methane (Hester et al., 2007). A more complex hydrate, composed of coexisting structure II and structure H hydrate, has been identified in a sample from the Barkley Canyon on the northern Cascadian margin. Surprisingly, the samples contain, beside methane and other

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. Figure 2 Variety of hydrate cavities; (a) Pentagonal dodecahedron 512 occupied with a methane molecule (b) Hexakaidecahedron 51264 occupied with a n-butane molecule (c) Combination of two Hexakaidecahedron 51264 occupied with n-butane and an iso-butane molecule respectively and a pentagonal dodecahedron 512 occupied with a methane molecule (modified from Luzi et al., 2008).

lighter hydrocarbons, some molecules which were not known as hydrate formers before, such as pentane and n-hexane. For the first time, these hydrate structures and compositions could be verified directly using powder X-ray diffraction and solid state 13C NMR (Lu et al., 2007).

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Hydrate Formation Processes

Hydrate formation can be specified as hydrate nucleation as the first step and hydrate growth as the second step. The nucleation process can be defined as a microscopic process involving tens of thousands molecules which first form small clusters and further develop into hydrate nuclei. By the time these small nuclei obtain a critical size, a continuous hydrate growth process starts. Three different hypotheses for hydrate nucleation are discussed here: the labile cluster nucleation hypothesis, the nucleation at the interface hypothesis and the local structuring hypothesis. The labile cluster nucleation hypothesis was first presented by Sloan and Fleyfel (1991) for the formation of hydrates from gas and ice and modified and extended by Christiansen and Sloan (1994): It starts with the presumption that the molecules of pure water without any dissolved gas molecules form labile ring structures of pentamers and hexamers. These labile water rings will construct labile clusters around gas molecules after gas is dissolved in the water phase. The coordination number of the water molecules depends on the size of the guest molecules, e.g., 20 water molecules surrounding one methane molecule, whereas 24 water molecules surround 1 ethane molecule and 28 water molecules cover 1 propane molecule. The clusters of the dissolved species combine to form unit cells. The formation rate of a particular

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hydrate structure depends on the availability of labile clusters with the required coordination numbers: From a mixture of methane and propane dissolved in liquid water hydrates should form more rapidly compared to either pure methane or propane dissolved in the water phase. The hypothesis of the nucleation at the interface was presented by Rodger (1990) and Kvamme (1996). It suggests the transportation of the gas molecules to the water–gas interface, where the gas molecules are adsorbed at the aqueous surface and diffuse to a suitable location. At this location the gas molecules will be enclathrated into the cavity, formed from water molecules. These cavities either agglomerate or grow by the addition of gas and water molecules into the vapor side of the interface. The local structuring nucleation hypothesis is based on the Landau free energy calculations performed by Radhakrishnan and Trout (2002). These calculations for carbon dioxide hydrate nucleation at the water–liquid carbon dioxide interface lead to the assumption that a group of CO2 molecules arrange in a configuration similar to that in the hydrate phase. This causes a local order of the surrounding water molecules which is different from that in the bulk water phase. If the number of CO2 molecules in this arrangement with a local order exceeds that of a critical hydrate nucleus, the formation of a hydrate nucleus starts. Regarding the hydrate growth three important aspects have to be considered: the transportation of gas and water molecules (mass transfer), the kinetics of the hydrate growth process and – due to the fact that hydrate formation is an exothermic process – the heat transfer away from the reaction (growth) site. The clathrate hydrate growth model presented by Englezos et al. (1987) is based on mass transfer theories. It describes the growth of the hydrate as a threestep process. The first step is the transport of the gas molecule into the liquid phase, the second step is the diffusion of the gas molecule through a stagnant liquid diffusion layer which surrounds the hydrate particle, and the last step is the incorporation of the gas molecule into the structured water framework of the hydrate particle, in the so-called ‘‘reaction’’ layer. Due to the fact that a concentration gradient of the gas molecules in the stagnant liquid layer is not allowed, the diffusion rate of the gas molecule through the stagnant liquid layer and the incorporation rate of the gas molecule into the hydrate structure are equal at steady state. Uchida et al. (1999) presented a model focusing on heat transfer which describes the formation of a hydrate film at the water–liquid carbon dioxide interface. They observed the primary nucleation at the interface and occasionally a secondary nucleation on the primary film. However, the propagation rate was temperature dependent which indicates that the heat diffusion is a restrictive factor. The model was recently modified and generalized by Mochizuki and Mori (2006) describing a heat-transfer-controlled lateral growth of a hydrate film at the interface between liquid water and an immiscible hydrate-forming fluid. Time resolved Raman spectroscopic measurements on methane hydrates during the formation process indicate that pentagonal dodecahedrons 512 of structure I are formed preferentially at the initial stages and that the formation of tetrakaidecahedrons 51262 may be the rate-limiting factor (Subramanian and Sloan, 1999; Uchida et al., 2000). Recently, time resolved Raman spectroscopic investigations on hydrates formed from ice and a gas mixture such as methane-H2S indicate a clear preference of the guest molecules regarding formation and occupancy of cavities. As shown in > Fig. 3, the Raman spectra indicate a preferred incorporation of H2S into tetrakaidecahedrons 51262 during the initial stages of hydrate formation whereas methane is preferentially incorporated into dodecahedrons 512 of structure I (Schicks et al., 2008). The observed cage occupancies with methane or H2S during the initial stages of hydrate formation do not correspond to those of the hydrate phase at equilibrium conditions.

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. Figure 3 Above: Real-time Raman spectra monitoring the incorporation of H2S in large 51262 (2,592 cm 1) and small 512 (2,602 cm 1) cavities of structure I H2S-CH4-hydrate. Below: Real-time Raman spectra monitoring the incorporation of CH4 in large 51262 (2,903 cm 1) and small 512 (2,915 cm 1) cavities of a mixed structure I H2S-CH4-hydrate.

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Regarding gas hydrate formation in nature it is questionable if a hydrate-forming fluid phase – e.g., a free gas phase – is available. In case of the absence of a free gas phase the formation of gas hydrates is limited to the availability of dissolved gas molecules. Furthermore, the presence of sediments and their influence on the hydrate formation process has to be considered when modeling hydrate formation processes. For the estimation of stabilization effects of gas hydrates on continental slopes, it is necessary to know how and where hydrate forms in the sediments. In the case of hydrate particle formation in contact with sediment grains, the hydrate phase may interact as cement and a hydrate saturation of less than 3% may already results in a stabilizing effect (Bernabe´ et al., 1992). Recent studies showed that the formation process of hydrates in sediments strongly depends on the water content and the absence or presence of a free gas phase. Klapproth et al. (2007) showed that hydrates between sediment (quartz) grains behave like cement in the presence of a free gas phase and with a water content of 10–17 wt%. However, other authors describe hydrate growth without any contact with the sediment grains when the water content is high both in presence of a free gas phase (Tohidi et al., 2001) and in absence of a free gas phase (Schicks et al., 2007).

4

Thermodynamic Properties and Hydrate Formation Kinetics

The thermodynamic properties of a hydrate phase describe the phase behavior including stability fields and decomposition conditions as well as metastable states and transition processes. In particular the stability field of a hydrate phase depends on its composition. Gas hydrates, containing nitrogen beside methane are less stable than pure methane hydrates: the stability field of these hydrates is shifted to higher pressures and lower temperatures compared to the stability conditions of pure methane hydrates (Jhaveri and Robinson, 1965). In contrast, mixed gas hydrates containing H2S, CO2 or higher hydrocarbons such as propane beside methane, are more stable than pure methane hydrates. Experimental data presented in > Fig. 4 show that the stability fields of these mixed hydrates are shifted to lower pressures and higher temperatures. The experimental data are in good agreement with calculated data (e.g., using CSMGem). Investigations on mixed gas hydrates containing small amounts of C3H8 (less than 5 vol% C3H8 beside methane in the feed gas), reveal an interesting phase behavior in the course of a transformation process as p-T-conditions approach the decomposition line of pure methane (Schicks et al., 2006). During this transformation process a simultaneous formation and decomposition of hydrate crystals was observed. Additionally, the morphology of the system changed from larger euhedral crystals to a foamy fine crystal mass. Raman spectroscopic and X-ray diffraction measurements indicate that the formation and decomposition of structure I hydrate in coexistence with structure II CH4-C3H8-hydrate are characteristic. The process could be observed by passing over the defined p-T-conditions (transformation conditions) in all directions. The reversibility indicates rather a thermodynamically stable phenomenon, than a metastable event. The lifetime of this transformation behavior – rapid crystal formation and decomposition – could be detected as a function of the sub-cooling below the transformation line: when within0.2 K of the transformation temperature at a given pressure, this transformation process could be observed for hours. The transformation process could not be clarified on a molecular level yet. However, the experimental data indicate that under defined pressure and temperature conditions the coexistence of several hydrate phases with different structure

Natural Gas Hydrates

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. Figure 4 P-T-diagrams based on experimental data presenting the stability fields of methane hydrate and mixed hydrates. The diagram above shows the stability fields for pure methane hydrate versus CO2–CH4 and H2S–CH4 hydrates. The diagram below shows the stability fields of pure methane hydrate versus CH4–C3H8 and CH4–C2H6–C3H8–hydrates. Independent from structure I or structure II formers, the decomposition conditions for the mixed hydrates are shifted to lower pressures and higher temperatures compared to pure methane hydrate.

and/or composition is favorable for the system from an energetic point of view. As already mentioned, a similar coexistence could also be proofed in nature e.g., in gas hydrate samples from Cascadia margin with coexisting structure II and structure H hydrate phases (Lu et al., 2007).

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Research Needs

Before gas hydrates were discovered in natural environments, they were recognized as a problem in natural gas pipelines due to the formation of undesirable hydrate plugs. Therefore laboratory investigations on hydrate formation and decomposition conditions were performed before the first hydrate deposit in the Messoyakha field was discovered in 1967 (Sloan, 1998). However, the results of laboratory and field studies are documented in a huge amount of papers. Nevertheless, there are still open questions in many areas of gas hydrate research, beginning at molecular level with the understanding of hydrate nucleation and growth processes. Also, as already mentioned earlier, the influence of hydrates on sediments and vice versa is not sufficiently clarified yet. In addition, innovative concepts for more applied topics, such as the improvement of gas hydrate detection in natural environments or the extraction of methane from natural gas hydrates are potential tasks for future research.

References Barnes RO, Goldberg ED (1976) Methane production and consumption in anoxic marine sediments. Geology 4: 297–300. Bernabe´ Y, Fryer DT, Hayes JA (1992) The effect of cements on the strength of granular rocks. Geophys Res Lett 19: 1511–1514. Bernard BB, Brooks JM, Sackett WM (1976) Natural gas seepage in the Gulf of Mexico. Earth Planet Sci Lett 31: 48–54. Boetius A, Ravenschlag K, Schubert CJ, Rickert D, Widdel F, Giesecke A, Amman R, Jørgensen BB, Witte U, Pfannkuche O (2000) A marine microbial consortium apparently mediating anaerobic oxidation of methane. Nature 407: 623–626. Cherskiy NV, Tsarev VP, Nikitin SP (1985) Investigations and predictions of conditions of accumulation of gas resources in gas-hydrate pools. Pet Geol 21: 65–89. Christiansen RL, Sloan ED (1994) Mechanisms and Kinetics of Hydrate Formation. In International Conference on Natural Gas Hydrates, New York Academy of Science, vol. 715. ED Sloan, J Happel, and MA Hnatow (eds.). New York: New Paltz, pp. 283–305. Dallimore SR, Uchida T, Collett TS (1999) Scientific results from JAPAX/JNOC/GSC Mallik 2 L-38 gas hydrate research well, Mackenzie Delta, Northwest Territories, Canada. Geol Surv Can Bull 544: 295–311. Englezos P, Kalogerakis N, Dholabhai PD, Bishnoi PR (1987) Kinetics of formation of methane and ethane gas hydrates. Chem Eng Sci 42: 2647–2658. Hester KC, Dunk RM, White SN, Brewer PG, Peltzer ET, Sloan ED (2007) Gas hydrate measurements at Hydrate Ridge using Raman spectroscopy. Geochim Cosmochim Acta 71: 2947–2959. Jhaveri J, Robinson DB (1965) Hydrates in the Methane– Nitrogen System. Can J Chem Eng 43: 75–78.

Kastner M, Kvenvolden K A, Lorenson TD (1998) Chemistry, isotopic composition, and origin of a methane-hydrogen sulfide hydrate at the Cascadia subduction zone. Earth Planet Sci Lett 156: 173–183. Klapproth A, Techmer KS, Klapp SA, Murshed MM, Kuhs WF (2007) Microstructure of gas hydrates in porous media. In Physics and Chemistry of Ice. WF Kuhs (ed.). Cambridge: RSC Publishing, pp. 321–328. Kvamme B (1996) A new theorie for kinetics of hydrate formation. In Proceedings of the 2nd International Conference on Natural Gas Hydrates, 2–6 June, Toulouse, France, pp. 139–146. Kvenvolden KA (1995) A review of the geochemistry of methane in natural gas hydrate. Org Geochem 23: 997–1008. Kvenvolden KA, Lorenson TD (2001) The global occurrence of natural gas hydrate. In Natural Gas Hydrates: Occurrence, Distribution, and Detection. CK Paull and WP Dillon (ed.). Washington: American Geophysical Union. Lu H, Seo Y, Lee J, Moudrakovski I, Ripmeester JA, Chapman NR, Coffin RB, Gardner G, Pohlman J (2007) Complex gas hydrate from Cascadia margin. Nature 445: 303–306. Luzi M, Schicks JM, Naumann R, Erzinger j, Udachin K, Moudrakowski I, Ripmeester JA, Ludwig R (2008) Investigations on the influence of guest molecule characteristics and the presence of multicomponent gas mixtures on gas hydrate properties In Proceedings of the 6th International Conference on Gas Hydrates, Vancouver. Mak TCW, McMullan RK (1965) Polyhedral clathrate hydrates. X: Structure of the double hydrate of tetrahydrofuran and hydrogen sulfide. J Chem Phys 42: 2732–2737.

Natural Gas Hydrates McMullan RK, Jeffrey GA (1965) Polyhedral clathrate hydrates. IX: Structure of ethylene oxide hydrate. J Chem Phys 42: 2725–2732. Milkov AV (2003) Global estimates of hydrate-bound gas in marine sediments: How much is really out there? Earth-Sci Rev 66: 183–197. Mochizuki T, Mori YH (2006) Clathrate-hydrate film growth along water/hydrate-former phase boundaries – numerical heat-transfer study. J Cryst Growth 290: 642–652. Radhakrishnan R, Trout BL (2002) A new approach for studying nucleation phenomena using molecular simulations: Application to CO2 hydrate clathrates. J Phys Chem 117: 1786–1796. Ripmeester JA, Tse JS, Ratcliffe CI, Powell BM (1987) A new clathrate hydrate structure. Nature 325: 135–136. Rodger PM (1990) Stability of gas hydrates. J Phys Chem 94: 6080–6089. Schicks JM, Naumann R, Erzinger J, Hester K, Koh CA, Sloan ED (2006) Phase Transitions in mixed gas hydrates: Experimental observations versus calculated data. J Phys Chem B 110: 11468–11474. Schicks JM, Luzi M, Erzinger J, Spangenberg E (2007) Clathrate hydrate formation and growth: Experimental data versus predicted behaviour. In Physics and Chemistry of Ice. WF Kuhs (ed.) Cambridge: RSC Publishing, pp. 537–544. Schicks JM, Luzi M, Spangenberg E, Nauman (See chapter 12 Volume 1, Marine cold seeps)n R, Erzinger J (2008) Hydrate formation investigations and kinetic studies under various defined conditions. In Proceedings of the 6th International Conference on Gas Hydrates, Vancouver. Schoell M (1988) Multiple origins of methane in the earth. Chem Geol 71: 1–10. Sloan ED Jr. (1998) Clathrate Hydrates of Natural Gases. New York: Marcel Dekker Inc.

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Sloan ED Jr. Fleyfel F (1991) A molecular mechanism for gas hydrate nucleation from ice. AIChE 37: 1281–1292. Sloan ED Jr., Koh CA (2008) Clathrate Hydrates of Natural Gases, 3rd edn. Boca Raton: CRC Press, Tayler and Francis Group. Subramanian S, Sloan ED (1999) Molecular measurements of methane hydrate formation. Fluid Phase Equilib 158–160: 813–820. Subramanian S, Kini RA, Dec SF, Sloan ED (2000) Evidence of structure II hydrate formation from ethane +methane mixtures. Chem Eng Sci 55: 1981–1999. Suess E, Torres ME, Bohrmann G, Collier RW, Rickert D, Goldfinger C, Linke P, Heuser A, Sahling H, Heeschen K, Jung C, Nakamura K, Greinert J, Pfannkuche O, Trehu A, Klinkhammer G, Whiticar MJ, Eisenhauer A, Teichert B, Elvert M (2001) Sea floor methane hydrates at hydrate ridge, Cascadia Margin. In Natural Gas Hydrates: Occurrence, Distribution, and Detection. CK Paull and WP Dillon (ed.). Washington: American Geophysical Union. Tohidi B, Anderson R, Clennell MB, Burgass RW, Biderkab AB (2001) Visual observation of gashydrate formation and dissociation in synthetic porous media by means of glass micromodels. Geology 29: 867–870. Uchida T, Ebinuma T, Kawabata J, Narita H (1999) Microscopic observations of formation processes of clathrate-hydrate films at an interface between water and carbon dioxide. J Cryst Growth 204: 348–356. Uchida T, Okabe R, Mae S, Ebinuma T, Narita H (2000) In situ observation of methane hydrate formation mechanism by Raman spectroscopy. Ann N Y Acad Sci 912: 593–601. Zehnder AJB, Brock TD (1979) Methane formation and methane oxidation by methanogenic bacteria. J Bacteriol 137: 420–432.

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4 Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes: Analytical Methods, Diversity, Characteristics W.-R. Abraham Helmholtz Center for Infection Research, Chemical Microbiology, Braunschweig, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 80

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Distillation and Extraction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81

3 3.1 3.2 3.3

Separation and Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Gas Chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Liquid Chromatography . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84 Mass Spectrometry as an Additional Dimension of Separation of Closely Related Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 84 3.4 Structure Elucidation and Isotopic Ratio . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 85 4

Terpenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

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Fats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87

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Hydrocarbons and Waxes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 89

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Ecological Roles of these Secondary Metabolites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 93

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_4, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: In the last century water distillation, the method of choice for many centuries to extract hydrophilic compounds from complex biological matrices, was more and more replaced by various chromatographic methods. At the same time chemical degradation steps to elucidate the structure of the compounds were replaced by physical methods. The current development goes to hyphenated instruments where multi-dimensional separations are combined with the subsequent structure elucidation of the individual compounds. The considerable improvements achieved in the analytical field led to the discovery and characterization of compounds never recognized before and allowed new insights in chemical ecology. Although essential oils, fats and waxes have been used for thousands of years, their main compositions have been revealed only in the last two centuries. Today more than 50,000 terpenes and about the same number of hydrocarbons are known. Since more sophisticated analytical instruments with higher sensitivities and resolutions become available each year their numbers are still increasing. With the growing numbers of compounds available new applications in various fields are realized. Although still in its infancy, this also includes a deeper understanding of the ecological functions of these hydrophobic compounds, which in itself presents many novel applications.

1

Introduction

Over the last two centuries more and more sophisticated methods became available to analyze complex organic mixtures like oil, food extracts or body liquids (> Fig. 1). In this short review an attempt is made to give an overview over this increasingly complex field. To achieve this mainly review articles have been cited to offer the reader a quick in-depth insight into the corresponding fields. Although we possess today a fleet of hyphenated instruments the old experience is still true that every new technique applied in a given field has the potential to uncover compounds and characteristics of compounds never recognized before. Oils, fats and waxes are usually defined by their physical properties which fit neither to their chemical classification nor to their biosynthesis. To keep things in line the different compound classes are defined first following their biosyntheses: Isoprenoids are natural products synthesized via mevalolactone or 1-desoxy-D-xylulose and consist of isoprene units. The nomenclature is that a single isoprenyl moiety is called a hemiterpene, two are a monoterpene, three a sesquiterpene, four a diterpene, five a sesterpene, six a triterpene and eight a tetraterpene or carotenoid. However, everything which follows the so called ‘‘isoprene rule’’ is not always a terpene as has been shown in the case of the fusalanipyrone (Abraham et al., 1990) or some terpenes are so heavily rearranged that it is hard to see the terpene in the final metabolite, e.g., in the cytotoxic mero-terpene sampsonione J (Hu and Sim, 1999) (> Fig. 2). Almost all essential oils produced by steam distillation of plant material consist of terpenes, mainly mono- and sesquiterpenes. Sterols are triterpenes which lost one or more carbons, so called nor-terpenes and steroids formed via cholesterol and having many functions as hormones in mammals lost the side chain almost completely. Fats are esters of long-chain fatty acids with a diol or triol, usually glycerol. If a long-chain fatty acid is esterified with a long-chain primary alcohol it will be defined as wax. Myricin is the ester between palmitic acid and myricylalcohol and the main component of beeswax. Long-chain hydrocarbons formed by the enzymatic decarboxylation of their corresponding fatty acids in plants and animals have also the physical properties of waxes, however, here they are defined as long-chain hydrocarbons.

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. Figure 1 Hydrophobic compounds are applied in many areas: From the seed of sunflowers oil is produced. Rosin (colophonium), refined from distilled oil of turpentine is applied by musicians to the hairs of the bow of a string instrument to increase the friction between bow and strings. Bees wax and long-chain hydrocarbons are burned in candles to give light but polycyclic aromatic hydrocarbons as produced as well.

Due to the difficulty to cover the entire diversity of hydrophobic natural products only examples will be given. In many ways the history of structure elucidation of natural products parallels the history of the development of the modern tools of analytical chemistry. As our ability to probe more precisely the exact nature of such molecules has advance, the true complexity of the substances present in oils and fats has become more apparent.

2

Distillation and Extraction

One of the oldest techniques to extract oils from biologic material is water distillation. Here water vapour is used to transport water non-mixable substances to cooled devices where the steam is condensed and the organic phase, the essential oil, is separated from the water. Water distillation is preferred against direct distillation because the thermic stress for the volatiles is much less allowing the survival of sensitive compounds. Water distillation is not a simple thing but an art and generations of essential oil producers developed it nearly to perfection. An alternative for water distillation is the extraction with all kinds of organic solvents. The polarity of the solvents controls the range of compounds extracted and the limitation here is only the need to have an organic phase which can easily be separated from the water phase. The choice for the solvent is limited by the necessity to separate it afterwards from the extracted oil. Because this is usually done by distillation low boiling solvents are preferred. A special position has here the extraction with supercritical gases (King, 2002). They are used under slightly elevated temperatures and high pressure and recycled by simply releasing the pressure causing the solvent to vaporize. Most common used is supercritical CO2 which is

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. Figure 2 Regular and irregular mono-, sesqui- and diterpenes discussed in the text.

rather non-polar and can be fine tuned in its polarity by the addition of small amounts of modulators, e.g., methanol. An alternative is solid phase extraction (SPE) where a solid is used to absorb the organic compounds. For more than a century SPE was limited to silica or aluminium oxide which are polar solids and not very selective. The advance in the modification of silica allowing the molecular binding of a very broad spectrum of organic residues to its surface together with the advances in inorganic chemistry leading to a fine-tuning in the structure and size of silica

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opened a completely new era in SPE. Nowadays alkyl chains from C2 to C30 with or without polar head groups like NH2 or OH offer the possibility to select solid phase adsorbents very specifically for the intended extraction protocol (Panagiotopoulou and Tsimidou, 2002). Many companies offer ready-to-use kits for the extraction of specific hydrophobic compounds, e.g., polycyclic aromatic hydrocarbons, polychlorinated diphenyls, herbicides, etc. To overcome the problem of small sample sizes together with a high dilution of the wanted compound miniaturizations of the extraction devices have been developed. One of the most important achievements here is the establishment of solid phase micro-extractions (SPME). SPME is based on a fused-silica fibre, coated with a polymeric stationary phase and mounted on a modified GC syringe that is exposed to the sample matrix. Because equilibrium is often not achieved a tight protocol is needed to ensure reproducibility (Dietz et al., 2006). To allow automation of SPME under the condition of high reproducibility solid phase dynamic extraction (SPDE) has been established in the last years (Bicchi et al., 2004). It utilizes a syringe attached to a stainless steel needle that is coated on its inner side with an immobilized extraction phase. For the extraction, the needle can be immersed directly into the aqueous sample or in the headspace. The syringe plunger is moved up and down several times for extraction, and the analytes are adsorbed in the internal coating. After several extraction cycles, the analytes are thermally desorbed from the coating in a GC injector. Compared with a SPME fibre, SPDE needle coatings possess up to six times larger extraction phase volumes and the whole device has the advantage of reduced fragility of the needle part, and the higher extraction speed (Demeestere et al., 2007).

3

Separation and Detection

Natural products usually reside in matrices and are often notoriously complex. Separation of its constituents is almost always needed to answer the research or industrial analysis questions which may be for comparative purposes for quality control, to discover new components, or to characterise the chemical classes of compounds present. The method of choice is today chromatography as the provider of separation often in combination with mass spectrometry to aid identification.

3.1

Gas Chromatography

Gas chromatography (GC) is still the method of choice for volatile compounds. Here the sample is evaporated in the injection port and transported by a gas stream through a capillary column where the compounds are separated. The detector most often used for hydrocarbons and essential oils is a flame ionization detector (FID) (Marriot et al., 2001). The diversity of GC column available together with tailor-made temperature programs allowed highly sophisticated applications in many fields, e.g., separation of mycolic acids (Butler and Guthertz, 2001). A special case is presented by archaeal ether lipids. These ether lipids are first hydrolysed and the ether lipids are then cleaved by hydrogen iodide and finally reduced to the hydrocarbons. Analysis of these hydrocarbons is achieved by GC often in connection with a mass spectrometer (Koga and Morii, 2006). The increasing need for higher resolution to analyze more complex mixtures was initially met by increasing the length of the GC-column because the resolution can be doubled for a specific compound pair if the column length is extended four times. However, e.g., a column length of 400 m and retention times of up to 11 h could

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not achieve the total separation of fuel samples (Berger, 1996). A more efficient way of resolution enhancement is through multidimensional GC where selected bands of overlapping compounds are transferred to a second column and separated by optimized conditions (Eyres et al., 2006). The revolutionary aspect of comprehensive two-dimensional gas chromatography (GC  GC), with respect to classical multidimensional chromatography, is that the entire sample is subjected to two distinct analytical separations. The resulting enhanced separating capacity makes this approach a prime choice for the separation of highly complex mixtures as demonstrated in the field of oil analyses (Blomberg et al., 2002). The addition of a third mass spectrometric dimension to a GC  GC system generates the most powerful analytical tool today for volatile and semi-volatile analytes (Mondello et al., 2008). Atmospheric pressure chemical ionization mass spectrometry (APCI-MS) has proven to be a very valuable technique readily producing useful ions with gentle fragmentation from large neutral molecules such as triacylglycerols and carotenoids, which are often difficult to analyze using other techniques (Byrdwell, 2001; Rezanka and Sigler, 2007).

3.2

Liquid Chromatography

Many compounds are simply not volatile enough to qualify for GC analysis. For many essential oil compounds it has been found that volatile compounds are generated from conjugates destroyed by the extraction process and a number of protocols have been established to analyse and quantify the intact conjugates (Moreau et al., 2002). Thin layer chromatography (TLC) has still its niche in certain applications (Sherma and Fried, 2005) but it is more and more replaced by all variations of high-performance liquid chromatography (HPLC). Some nonpolar compounds like polyyne hydrocarbons and highly unsaturated fatty acids are too unstable for GC-analyses and have to been separated by TLC or HPLC. To improve the separation of these often very similar compounds the TLC plate or HPLC column have been impregnated with silver ions. The metal ion interacts with the double or triple bonds and retards the migration of highly unsaturated metabolites (Ratnayake, 2004). HPLC has its advantage for mixtures with UV-absorbing compounds. The analysis of carotenoids in plasma, foods and tissues has thus become of interest in studies examining the role of diet in chronic disease prevention because epidemiologic and clinical studies have shown that a high intake of vegetables and fruit, with consequently high intakes and circulating concentrations of carotenoids, is associated with reduced risk of cardiovascular and other chronic diseases. High-performance liquid chromatography with ultra-violet or photodiode array detection is most often employed in routine use (Su et al., 2002). For compounds lacking such chromophores a number of detectors have been developed based e.g., on changes in the refraction index, the dielectric constant or the thermal conductivity of the effluent (Brondz, 2002). Only the light scattering detector found a niche for certain application but is rapidly replaced by mass selective detectors or mass spectrometers.

3.3

Mass Spectrometry as an Additional Dimension of Separation of Closely Related Compounds

Until now mass spectrometry has only been discussed as means to detect an eluting compound but mass spectrometry offers much more than detection and peak identification. Many

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different mass spectrometric techniques are currently available to analyse and identify mixtures of compounds. Although these instruments are rarely used as stand-alone instruments but usually connected to a GC or HPLC system they are discussed separately here to account for their complexity. One of the simplest methods to identify compounds is a mass spectrometer connected to a GC. Either a given compound is identified by comparison with the spectra library of the software or the compound produces characteristic fragments in the mass spectrometer which facilitates identification. An ion-trap or multiple MS (MSn) allows the analysis of any molecular ion in a complex mixture or fragment ion of interest, e.g., by its decay by collision-induced-dissociation (CID). The introduction of the desorption–ionization techniques of fast atom bombardment (FAB) and electrospray (ES) circumvented the problem of analyte volatility and thermal stability (Pulfer and Murphy, 2003), whereas HPLC can be directly coupled with these ionization methods to provide chromatographic separation. FAB and ES both tend to give only molecular weight information, but to enhance structural information, tandem mass spectrometry (MS/MS) experiments can be performed. In an MS/MS experiment, a precursor ion is initially selected by the first mass analyzer (MS1) and induced to fragment by collision with an inert gas. The resultant fragment ions are analyzed by a second mass analyzer (MS2) and the structure of the precursor deduced from the resultant product-ion spectrum (Griffiths, 2003). Quadrupole–time-of-flight (TOF) mass spectrometers combine the high performance of time-of-flight analysis in both the mass spectrometry (MS) and tandem MS (MS/MS) modes, with the widely used techniques of electrospray ionization (ESI) and atmospheric pressure chemical ionization (APCI) (Chernushevich et al., 2001). The determination of unknown structures is usually time-consuming, however, the highperformance features of high magnetic field Fourier Transform Ion Cyclotron Resonance Mass Spectrometry (FTICR-MS) have greatly alleviated the structural elucidation bottleneck. The high-performance features of high field FTMS include unsurpassed mass measurement accuracy for elemental formula determination, ultra-high mass resolution for component separation, the ability to perform multiple levels of tandem mass spectrometry for structural elucidation, and moderate sensitivity for limited supply of isolates (Feng and Siegel, 2007). The high price is currently the most important drawback for the broad application of such instruments.

3.4

Structure Elucidation and Isotopic Ratio

To elucidate the structure of a natural compound usually requires the combination of different techniques, often infrared (IR) and ultraviolet (UV) spectroscopy, mass spectrometry (MS) and nuclear magnetic resonance (NMR) spectrometry. Each technique allows a number of variations also in hyphenation with GC and HPLC to fit the specific requirements for the unknown compound and a number of textbooks have been published especially on MS and NMR applications. The majority of natural products are asymmetric due to chiral centers. The absolute configuration of a compound is often essential for its biological activity, only to mention here the caraway-like smelling S(+)- and spearmint-like smelling R( )-carvone (Friedman and Miller, 1971) and nootkatone where the 4R,4aS,6R-(+)-enantiomer has a grapefruit odor while the (+)-isomer is woody and spicy (Bentley, 2006) (> Fig. 3). To assess the absolute

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. Figure 3 Examples for bioactive enantiomers.

configuration of natural compounds a number of techniques have been applied. The simplest method is the derivatization with an enantiomerically pure chiral reagent leading to diastereomers. The Mosher ester method using the esterification with both of the optically pure alpha-methoxy-alpha-trifluoromethylphenylacetic acids enables the direct determination of the absolute configuration by characteristic shift differences observed in 1H-NMR (Hoye et al., 2007). GC- or HPLC-columns loaded with an enantiomerically pure compound (e.g., cyclodextrin (Bicchi et al., 1999) or amino acid derivative) have been used to separate the enantiomers and chiral shift reagents have been applied in NMR. Where the pure enantiomer is available, these techniques allow the determination of the absolute stereochemistry and the enantiomeric excess in the mixture analysed (Gu¨bitz and Schmid, 2006). Enantioselectivity as well as isotope discrimination during biosynthesis, may serve as ‘‘endogenous’’ parameters, provided that suitable methods and comprehensive data from authentic sources are available. Enantioselective capillary gas chromatography and online methods of isotope-ratio mass spectrometry in the authentication of food flavor and essential oil compounds are currently here the methods of choice (Mosandl, 2004). Compound-specific carbon-isotope (13C/12C) ratios also enable quantitative assessment of sources of polycyclic aromatic hydrocarbons in natural environments and have been applied to source apportionment studies of PAHs in soils and sediments from marine, lacustrine, and terrestrial environments (Poster et al., 2006).

4

Terpenes

Terpenes are produced by organisms from all kingdoms, however, plants and fungi seem to have the highest diversity in this class of natural compounds with more than 25,000 compounds currently known (Dictionary of natural compounds, 2008). To have a feeling about the importance of terpene biosynthesis it should be noted that 500 Tg C a 1, about one third

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of the total volatile organic carbon emission on Earth, belongs to isoprene (Goldstein and Galbally, 2007). Monoterpene hydrocarbons are important ingredients of plants, resins and fruit peels. They function here as insecticides, antimicrobial agents and in wound sealing (Gershenzon and Dudareva, 2007). Representative monoterpenes are limonene, 3-carene and a-pinene, representative sesquiterpene alcohols are a-santalol and a-bisabolol and representative diterpene acids are abietic acid and pimaric acid used in many pharmaceutical applications (> Fig. 2). Due to their biosyntheses the carbon skeletons of terpenes are oligomers of isoprene but many of them undergo further modifications. An example for this is geosmin, responsible for the characteristic odor of moist soil and unpleasant off-flavors in water, which is produced by a number of microorganisms, including most Streptomyces and some fungi. Geosmin is one of the most intensive odours known, detectable for humans already in dilutions of 1 ppt in water. Its biosynthesis both in bacteria and fungi runs via cyclization of farnesyl diphosphate to germacradienol, fragmentation with a loss of acetone to 8,10-dimethyl-1-octalin and rearrangement the 1(9)-octalin and addition of water to the trinor-sesquiterpene geosmin (Jiang and Cane, 2008). Fungi became a rich source of structurally very diverse terpenes and a number of terpene skeletons seem to be confined to this kingdom. Several fungal sesquiterpenes, e.g., of the marasmane, hirsutane, fomannosane or fumigallane type possess antibiotic, cytotoxic or immunosuppressant activities which make them attractive as pharmaceutical lead structures (Abraham, 2001). The trichothecane sesquiterpenes, mainly produced by Fusarium spp., harbour numerous toxic compounds with trichothecin and crotocin as one of the less polar compounds (Kimura et al., 2007). The acyclic triterpene squalene is the key precursor for most tetra- and pentacyclic triterpenes, e.g., lanosterol. Many methyl groups of this triterpene are then cleaved off leading, e.g., to cholesterol. Side-chain methylated sterols are the phytosterols, ergosterol and sitosterol (> Fig. 4). Partial degradation of the side chain gives the class of steroids, among them many important human hormones. When Berzelius named yellow pigments from autumn leaves as xanthophylls in 1837 he was the first to realize this group of naturally occurring oxygenated carotenoids synthesized mainly by plants and microorganisms. While more than 300 different carotenoids and xanthophylls are currently known the hydrocarbons a- and b-carotene and the acyclic lycopene are the most important tetraterpenes in human nutrition. The enzymatic cleavage of the central double bond of b-carotene leads to retinol, also known as vitamin A1. Xanthophylls like lutein or zeaxanthin (> Fig. 5) have applications as natural feed additives, and as an active ingredient in medicinal pharmaceuticals due to their outstanding antioxidant properties. Dietary xanthophylls may inhibit the onset of many diseases such as arteriosclerosis, cataracts, age-related macular degeneration, multiple sclerosis, and cancers.

5

Fats

Fats or lipids can be divided into three main classes: Neutral lipids, glycolipids and phospholipids. Neutral lipids which are triesters of glycerol where the carboxylic acids are usually longchain fatty acids. Glycolipids are also esters of glycerol but here only two hydroxyl groups are esterified with long chain fatty acids, the third one forms an ether link to a sugar moiety.

87

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Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes

. Figure 4 Triterpenes and sterols.

Phospholipids are the most polar compounds in this group because the diglyceride is here esterified with phosphoric ester and usually further connected to another functional group. These categories are not completely separated because some phospholipids carrying sugar moieties and glycolipids with sulfonic acids or even phospolipds are also known. In Archaea the diglyceride moiety is replaced by terpene ethers usually of the phytane (sn-2,3-diphytanylglyceryl) or biphytanyl type. Cyclopentyl or cyclohexyl rings within the terpene side chain lead to a higher rigidity of the lipid (DeLong et al., 1998) important for creatures thriving at high temperatures. Neutral lipids are used by plants and animals as storage compounds. The ability of some plants to store large amounts of carbon and energy in neutral lipids led to their cultivation all over this planet first for nutrition but later on also for various chemical applications (> Fig. 1). Palm oil produced from the fruit of palms is today the main oil source worldwide but the demand for renewable energy led to the rapid increase of the production of rapeseed and other oils to be converted to bio-diesel. Long-chain fatty acids are usually esterified to glycerol but in minor amounts they also occur as free acids. Typical fatty acids of bacteria have 12–18 carbons and no or one double bond. Some of these fatty acids bear a methyl group at o-1- (iso) or o-2-position (anteiso) and

Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes

4

. Figure 5 Some of the most important carotenoids.

some sulphate reducing bacteria have it at C-10. Longer fatty acids with more double bonds are known from plants and fungi but have also been reported from Shewanella species (Bowman et al., 1997). Highly modified fatty acids have been reported as well, e.g., a hexamethano-octadecadien amide U-106305 from Streptomyces sp., an inhibitor of cholesteryl ester transfer protein (Kuo et al., 1995), acetylenic or cumulenic brominated fatty acids from lichens or leukotriene A4, a second messenger precursor of inflammations in humans (> Fig. 6). The South African plant genus Dichapetalum protects their leaves by o-fluorinated fatty acids including fluoroacetate acting as inhibitors in the citrate cycle of potential grazers (Dembitsky and Srebnik, 2002).

6

Hydrocarbons and Waxes

Crude oils are formed by physico-chemical modifications of biomass over millions of years. The diversity of compounds present in crude oil is impressive comprising beside all kinds of hydrocarbons hetero-aromatic compounds, ethers and ketones (Berger, 1996). Many organisms protect their outer surfaces with waxes. Usually these waxes comprise several chemical compound classes like long-chain hydrocarbons, alcohols, acids and their esters. Waxes are very spectacular at plants living under arid conditions like some Tillandsia (Bromeliaceae) or Copiapoa (Cactaceae) species which obscure the chlorophyll and give the body a white appearance. Less spectacular but equally important is the protection of insects with waxes against desiccation and microbial attacks (Juarez and Fernandez, 2007). The hydrocarbons in these mixtures are usually formed by decarboxylation of the corresponding long-chain fatty acids resulting in hydrocarbons with uneven numbers of carbons. Fatty acids, alcohols or hydrocarbons may also be methylated and branched hydrocarbons bearing up to 4 methyl groups are known.

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. Figure 6 Examples of fatty acids and derived bioactive secondary metabolites.

A special situation is found in insects using waxes as building material for their houses. The most famous example is the honey bee whose waxes are used for thousand of years by humans. More than 250 different compounds have been detected in beeswax and main constituents are myricin (65%), the ester between palmitic acid (CH3(CH2)14COOH) and myricyl alcohol (triacontanol, CH3(CH2)30OH), the ester between cerotic acid (CH3(CH2)24COOH) and triacontanol and a variety of long-chain alkanes like hentriacontane (CH3(CH2)29CH3). Mycobacterium and related genera produce ether-soluble ‘‘waxes,’’ known as mycolic acids. What was once known as mycolic acid is now a broad family of over 500 closely related homologous chemical structures. All mycolic acids possess at least two chiral centers at the positions a and b to the carboxylic acid which have the R,R-erythro configuration. For all organisms producing these molecules extended ethylenic series, differing in molecular weight by exactly 28 amu, are entirely typical. Beside the basic b-hydroxy-a-alkyl branched structure mycolic acids contain diverse functionalities (> Fig. 7). The unsaturations and cyclopropanes occurring within a-mycolates may be either cis or trans, but when trans they always also possess an adjacent methyl group. Although cyclopropanation is observed only in pathogenic mycobacteria little is known about the physiological function of this modification. Mycolic acids have become one of the defining taxonomic characteristics of many species in genera such as Mycobacterium, Corynebacterium, Dietzia, Nocardia, Gordona, Rhodococcus and Tsukamurella. While Corynebacterium produces corynomycolic acids with typical sizes of

Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes

4

. Figure 7 The main types of mycolic acids.

30–36 carbons with an a-branch of about 16–18 carbons, Nocardia has nocardomycolates of approximately 50 carbons with 0–3 double bonds and Gordona has mycolates of approximately 60 carbons with 1–4 double bonds. Finally, Tsukamurella has 62–78 carbon mycolates with 20–22 carbon a-branch and up to six double bonds (Barry et al., 1998). Almost two centuries ago the first acetylenic natural product was identified in chamomille and especially Ferdinand Bohlmann pioneered the discovery of hundreds of polyynes from plants (Zdero and Bohlmann, 1990). Due to the reactivity of the triple bond many of these compounds possess diverse bioactivities (Dembitsky, 2006) and have been used in bioactive ethnobotanicals for centuries (> Fig. 8).

7

Ecological Roles of these Secondary Metabolites

It is highly likely that all secondary metabolites have certain functions for the producing organism but we just started to understand some of them (Harborne, 2001). In the plant kingdom it becomes more and more obvious that the communication between plants and animals is achieved by a high diversity of volatiles (Pichersky et al., 2006). A well studied chemical communication system can be found between plants and insects, where upon attack plants secrete volatiles which attract insectivores (Schnee et al., 2006). From the lima bean (Phaseolus lunatus) upon feeding of the spider mite Tetranchys urticae the homo-monoterpene (3E)-4,8-dimethyl-1,3,7-nonatriene and the homo-sesquiterpene

91

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Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes

. Figure 8 Bioactive polyynes from bacteria, fungi and plants.

Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes

4

(3E,7E)-4,8,12-trimethyl-l,3,7,11-tridecatetraene (> Fig. 1) are emitted. These kairomones attract predatory mites which then feed on the spider mite (Dicke et al., 1990). Formation of the homo-terpenes is not observed after simply injuring the plants but can be triggered by treatment of the undamaged leaves with b-glucosidase. These two homo-terpenes trigger similar plant-herbivore-insectivore interactions in other plants.

8

Research Needs

Because of the high potential of high-resolution chromatography and the huge increase in available hyphenated instruments the next step is the investigation of hydrophilic compounds in complex biological systems under field conditions. Commercially available field SPME or portable GC-MS instruments are first developments in this direction. These questions can now be addressed using multidimensional separations with integrated instruments for structure elucidation in combination with new techniques, e.g., isotope ratio mass spectrometry or FT-MS. Although the available sensitivity of the individual instruments is already impressive further improvements to lower the amount of sample material and to work with trace compounds in complex matrices are still urgently needed. The history of analytical chemistry tells us that any major improvement of sensitivity and resolution will lead to entirely novel compounds and processes. Compounds produced in traces or in very low concentrations in complex matrices still present a challenge for modern analytics but from experience we know that many of them have completely novel characteristics making their investigation worthwhile. Hyphenated techniques often offer completely new insights into the ecology of producing organisms and the biosyntheses of known compounds. Now we can start to ask the questions: In which communities and under which conditions are certain compounds or series of compounds formed? What are their ecological functions? Learning more about natural products in the environment will surely help us in understanding the resilience of communities, their adaptation to environmental stress and – at the end – this will open entirely new fields of applications of these compounds in a multitude of biotechnological applications.

References Abraham W-R (2001) Bioactive sesquiterpenes produced by fungi: are they useful for humans as well? Curr Med Chem 8: 583–606. Abraham W-R, Knoch I, Witte L (1990) Biosynthesis of the terpenoidic polyketide fusalanipyrone Phytochemistry. 29: 2877–2878. Barry III CE, Lee RE, Mdluli K, Sampson AE, Schroeder BG, Slayden RA, Yuan Y (1998) Mycolic acids: structure, biosynthesis and physiological functions. Prog Lipid Res 37: 143–179. Bentley R (2006) The nose as a stereochemist. Enantiomers and odor. Chem Rev 106: 4099–4112. Berger TA (1996) Separation of a gasoline on an open tubular column with 1.3 million effective plates. Chromatographia 42: 63–71.

Bhosale P, Bernstein PS (2005) Microbial xanthophylls. Appl Microbiol Biotechnol 68: 445–455. Bicchi C, Cordero C, Liberto E, Rubiolo P, Sgorbini B (2004) Automated headspace solid-phase dynamic extraction to analyse the volatile fraction of food matrices. J Chromatogr A 1024: 217–226. Bicchi C, D’Amato A, Rubiolo P (1999) Cyclodextrin derivatives as chiral selectors for direct gas chromatographic separation of enantiomers in the essential oil, aroma and flavour fields. J Chromatogr A 843: 99–121. Blomberg J, Schoenmakers PJ, Brinkman UAT (2002) Gas chromatographic methods for oil analysis. J Chromatogr A 972: 137–173.

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Bowman JP, McCammon SA, Nichols DS, Skerratt JH, Rea SM, Nichols PD, McMeekin TA (1997) Shewanella gelidimarina. sp. nov. and Shewanella frigidimarina sp. nov., novel Antarctic species with the ability to produce eicosapentaenoic acid (20:5 omega 3) and grow anaerobically by dissimilatory Fe(III) reduction. Int J Syst Bacteriol 47: 1040–1047. Brondz I (2002) Development of fatty acid analysis by high-performance liquid chromatography, gas chromatography, and related techniques. Anal Chim Acta 465: 1–37. Butler WR, Guthertz LS (2001) Mycolic acid analysis by high-performance liquid chromatography for identification of Mycobacterium. species. Clin Microbiol Rev 14: 704–726. Byrdwell WC (2001) Atmospheric pressure chemical ionization mass spectrometry for analysis of lipids. Lipids 36: 327–346. Chernushevich IV, Loboda AV, Thomson BA (2001) An introduction to quadrupole/time-of-flight mass spectrometry. J Mass Spectrom 36: 849–865. Delong EF, King LL, Massana R, Cittone H, Murray A, Schleper C, Wakeham SG (1998) Dibiphytanyl ether lipids in nonthermophilic Crenarchaeotes. Appl Environ Microbiol 64: 1133–1138. Dembitsky VM (2006) Anticancer activity of natural and synthetic acetylenic lipids. Lipids 41: 883–924. Dembitsky, VM, Srebnik M (2002) Natural halogenated fatty acids: their analogues and derivatives. Prog Lipid Res 41: 315–367. Demeestere K, Dewulf J, De Witte B, Van Langenhove H (2007) Sample preparation for the analysis of volatile organic compounds in air and water matrices. J Chromatogr A 1153: 130–144. Dicke M, van Beek TA, Posthumus MA, Ben Dom N, van Bokhoven H, de Groot AE (1990) Isolation and identification of volatile kairomone that affects acarine predator-prey interactions. Involvement of host plant in its production. J Chem Ecol 16: 381–396. Dictionary of Natural Compounds (2008) Chapman and Hall Chemical Database. Dietz C, Sanz J, Ca´mara C (2006) Recent developments in solid-phase microextraction coatings and related techniques. J Chromatogr A 1103: 183–192. Eyres G, Marriott PJ, Dufour JP (2006) The combination of gas chromatography-olfactometry and multidimensional gas chromatography for the characterisation of essential oils. J Chromatogr A 1150: 70–77. Feng X, Siegel MM (2007) FTICR-MS applications for the structure determination of natural products. Anal Bioanal Chem 389: 1341–1363. Friedman L, Miller GJ (1971) Odor incongruity and chirality. Science 172: 1044–1046.

Furr HC (2004) Analysis of retinoids and carotenoids: problems resolved and unsolved. J Nutr 134: 281S–285S. Gershenzon J, Dudareva N (2007) The function of terpene natural products in the natural world. Nat Chem Biol 3: 408–414. Goldstein AH, Galbally IE (2007) Known and unexplored organic constituents in the Earth’s atmosphere. Environ Sci Technol 41: 1514–1521. Griffiths WJ (2003) Tandem mass spectrometry in the study of fatty acids, bile acids, and steroids. Mass Spectrom Rev 22: 81–152. Gu¨bitz G, Schmid MG (2006) Chiral separation principles in chromatographic and electromigration techniques. Mol Biotechnol 32: 159–180. Harborne JB (2001) Twenty-five years of chemical ecology. Nat Prod Rep 18: 361–379. Hoye TR, Jeffrey CS, Shao F (2007) Mosher ester analysis for the determination of absolute configuration of stereogenic (chiral) carbinol carbons. Nat Protoc 2: 2451–2458. Hu LH, Sim KY (1999) Cytotoxic polyprenylated benzoylphloroglucinol derivatives with an unusual adamantyl skeleton from Hypericum sampsonii. (Guttiferae). Org Lett 1: 879–882. Jiang J, Cane DE (2008) Geosmin biosynthesis. Mechanism of the fragmentation-rearrangement in the conversion of germacradienol to geosmin. J Am Chem Soc 130: 428–429. Juarez MP, Fernandez GC (2007) Cuticular hydrocarbons of triatomines. Comp Biochem Physiol Mol Integr Physiol 147: 711–730. Kimura M, Tokai T, Takahashi-Ando N, Ohsato S, Fujimura M (2007) Molecular and genetic studies of Fusarium. Trichothecene biosynthesis: pathways, genes and evolution. Biosci Biotechnol Biochem 71: 2105–2123. King JW (2002) Supercritical fluid extraction: present status and prospects. Grasas y Aceites 53: 8–21. Koga Y, Morii H (2006) Special methods for the analysis of ether lipid structure and metabolism in archaea. Anal Biochem 348: 1–14. Kuo MS, Zielinski RJ, Cialdella JI, Marschke CK, Dupuis MJ, Li GP, Kloosterman DA, Spilman CH, Marshall VP (1995) Discovery, isolation, structure elucidation, and biosynthesis of U-106305, a cholesteryl ester transfer protein inhibitor from UC 11136. J Am Chem Soc 117: 10629–10634 Marriot PJ, Shellie R, Cornwell C (2001) Gas chromatographic technologies for the analysis of essential oils. J Chromatogr A 936: 1–22. Mondello L, Tranchida PQ, Dugo P, Dugo G (2008) Comprehensive two-dimensional gas chromtagraphymass spectrometry: a review. Mass Spetrom Rev 27: 101–124.

Biosynthetic Oils, Fats, Terpenes, Sterols, Waxes Moreau RA, Whitaker BD, Hicks KB (2002) Phytosterols, phytostanols, and their conjugates in foods: structural diversity, quantitative analysis, and health-promoting uses. Prog Lipid Res 41: 457–500. Mosandl A (2004) Authenticity assessment: a permanent challenge in food flavor and essential oil analysis. J Chromatogr Sci 42: 440–449. Panagiotopoulou PM, Tsimidou M (2002) Solid phase extraction: applications to the chromatographic analysis of vegetable oils and fats. Grasas y Aceites 53: 84–95. Pichersky E, Noel JP, Dudareva N (2006) Biosynthesis of plant volatiles: nature’s diversity and ingenuity. Science 311: 808–811. Poster DL, Schantz MM, Sander LC, Wise SA (2006) Analysis of polycyclic aromatic hydrocarbons (PAHs) in environmental samples: a critical review of gas chromatographic (GC) methods. Anal Bioanal Chem 386: 859–881. Pulfer M, Murphy RC (2003) Electrospray mass spectrometry of phospholipids. Mass Spectrom Rev 22: 332–364. Ratnayake WMN (2004) Overview of methods for the determination of trans fatty acids by gas

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chromatography, silver-ion thin-layer chromatography, silver-ion liquid chromatography, and gas chromatography/mass spectrometry. J AOAC Int 87: 523–539. Rezanka T, Sigler K (2007) The use of atmospheric pressure chemical ionization mass spectrometry with high performance liquid chromatography and other separation techniques for identification of triacylglycerols. Curr Anal Chem 3: 252–271. Schnee C, Ko¨llner TG, Held M, Turlings TCJ, Gershenzon J, Degenhardt J (2006) The products of a single maize sesquiterpene synthase form a volatile defense signal that attracts natural enemies of maize herbivores. Proc Natl Acad Sci USA 103: 1129–1134. Sherma J, Fried B (2005) Thin layer chromatographic analysis of biological samples. A review. J Liq Chromatogr Relat Technol 28: 2297–2314. Su Q, Rowley KG, Balazs ND (2002) Carotenoids: separation methods applicable to biological samples. J Chromatogr B Analyt Technol Biomed Life Sci 781: 393–418. Zdero C, Bohlmann F (1990) Systematics and evolution within the Compositae, seen with the eyes of a chemist. Plant Syst Evol 171: 1–14.

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Part 2

Formation and Location

5 Stable Isotopes in Understanding Origin and Degradation Processes of Petroleum A. Vieth . H. Wilkes Helmholtz Centre Potsdam (GFZ), Organic Geochemistry, Potsdam, Germany [email protected] [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100

2

Definitions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100

3 Stable Isotope Applications in Petroleum Geochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . 102 3.1 Petroleum Formation in Sedimentary Basins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 3.2 Alteration Processes in Petroleum Reservoirs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 4

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 108

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_5, # Springer-Verlag Berlin Heidelberg, 2010

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Stable Isotopes in Understanding Origin and Degradation Processes of Petroleum

Abstract: In this essay we provide a short introduction to and overview of the basics of stable isotope geochemistry and its common application in petroleum geochemistry. We identify the processes that are responsible for the carbon and hydrogen isotopic compositions of biological and geological organic matter and indicate the utility of stable isotopes in oil-source rock correlations. Stable isotope analyses are also exploited in the investigation of different alteration processes within oils and petroleum reservoirs. State of the art work is presented, and future research needs are identified.

1

Introduction

Processes controlling the molecular and isotopic composition of petroleum – and thus its physico-chemical properties and quality – are divided into two fundamental categories. Primary processes include everything influencing petroleum composition prior to the accumulation in a trap (reservoir); these are for example, the biological origin of the source organic matter, the depositional environment of the source rock and its thermal maturation as well as migration of petroleum fluids from the source rock to the trap. Secondary controls lead to an alteration of reservoired petroleum after accumulation in the trap; this includes (bio)chemical processes such as biodegradation and thermochemical sulfate reduction and physical processes such as water washing and evaporative fractionation. The evaluation of these processes is challenging as petroleum reservoirs typically have complex filling histories with, for example, alternating charging and biodegradation events. Within this essay some general information about stable isotopes is given (Section 1). Section 2.1 attempts to be suggestive of the complexity of processes related to petroleum generation that have an influence on the isotopic composition of crude oil and natural gas, and describes some applications. Section 2.2 discusses the role of alteration processes within reservoirs with a main focus on microbial activity in these environments. It is not within the scope of this essay to give a comprehensive overview on the detailed factors influencing the isotopic composition of petroleum. Excellent reviews on this topic can be found in the recent literature (Galimov 2006; Peters et al., 2005).

2

Definitions

For a more comprehensive introduction to stable isotope geochemistry and the analytical methods for determining relative isotope ratios, the interested reader is advised to consult one of the following textbooks: Clark and Fritz 1997; Hoefs 2004; Sharp 2006. With respect to hydrocarbons the following essay will be focused on the stable isotopes of carbon and hydrogen. The isotopes of a given element have the same number of protons and electrons but differ from each other by the number of neutrons in the nucleus. Most elements in the periodic table have two or more naturally occurring isotopes; carbon and hydrogen both have two stable isotopes (> Table 1). The mass differences, resulting from the difference in the number of neutrons, lead to differences in the chemical and physical properties of molecules with light and/or heavy isotopes. This causes partial separation of the light isotopes from the heavy isotopes during chemical reactions (isotope fractionation). Equilibrium chemical reactions (e.g., dissolution of CO2 in water) are accompanied by equilibrium isotope effects, whereas unidirectional

5

Stable Isotopes in Understanding Origin and Degradation Processes of Petroleum

reactions – such as biodegradation – quite often are accompanied by significant kinetic isotope effects. In biological systems usually reactions with the lighter isotopes are preferred, due to the lower energy that is required to break a bond within one molecule between two light isotopes, in comparison to the higher energy that is needed to break the bond between a light and a heavy isotope. The delta (d) notation has been introduced to express the relative differences in isotopic compositions (McKinney et al., 1950),   Rsample  Rstandard 0  1000; ð1Þ d½ =00 ¼ Rstandard where R is the ratio of the abundance of the heavy to the light isotope (within the sample and the international standard; see > Table 2). R is given by D/H and 13C/12C, respectively. d values are reported in per mil, or parts per thousands. A negative d value means that the ratio of the heavy to the light isotope is lower in the sample than it is in the standard, the sample is ‘‘depleted’’ in the heavy isotope in comparison to the relatively ‘‘enriched’’ standard. Biodegradation processes typically lead to enrichment of the molecules with the heavier isotope in the residual fraction of a substrate due to kinetic isotope effects of the first irreversible reaction in a degradation pathway. The relation between decrease in concentration and change in isotopic composition of the residual substrate can be described by the Rayleigh Equation (2) where F is the fraction of the hydrocarbon remaining (C/Ci), R is the isotopic composition of the hydrocarbon at a particular F and Ri is the initial isotopic composition. R ¼ F ða1Þ : Ri

ð2Þ

. Table 1 Isotopic abundances and relative atomic masses of carbon and hydrogen (data taken from Sharp 2006) Symbol

Atomic number

Mass number

Abundance (%)

H

1

1

99.985

D

1

2

0.015

C

6

12

98.89

C

6

13

1.11

Atomic weight 1.007825 2.0140 12.0 13.00335

. Table 2 Names and relative and absolute isotope ratios of international standards Name

Description

Ratio

Accepted value

d value

PDB

Belemnitella americana from Pee Dee Formation in USA

13

11237.3  2.9 (Craig 1957)

0.00

SMOW

Standard Mean Ocean Water

D/H

155.76  0.10 (Hagemann 1970)

0.00

C/12C

101

102

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Stable Isotopes in Understanding Origin and Degradation Processes of Petroleum

The isotope fractionation factor (a) relates the changes in the isotopic composition to changes in the concentration of the residual fraction during the transformation. It is quite common to use the enrichment factor e (3) instead of a. The Rayleigh Equation provides the opportunity to quantify the extent of biodegradation based on carbon or hydrogen isotope ratios and the appropriate isotope fractionation factor independent from concentration measurements. e ¼ ða  1Þ  100:

3

Stable Isotope Applications in Petroleum Geochemistry

3.1

Petroleum Formation in Sedimentary Basins

ð3Þ

A key task of petroleum geochemistry is the correct correlation of natural gas or crude oil accumulations to their respective source rock(s) in a given exploration area. Any approach used in oil- or gas-source correlation is based on the assumption that similarities exist between the molecular and/or isotopic composition of a petroleum fluid and its source. The discussion in this subsection will focus on geochemical processes that have to be considered when using the stable carbon or hydrogen isotopic composition of organic matter as a correlation tool, although typically multiple independent data types, both molecular and isotopic, are used in such studies. The most important control on the carbon isotopic composition of crude oils and natural gas is the carbon isotopic composition of the source organic matter that has been deposited in the sediment later forming a petroleum source rock. The isotopic composition of biogenic organic matter depends on four key elements, namely (1) the carbon source utilized; (2) isotope effects associated with assimilation of carbon by the producing organism; (3) isotope effects associated with metabolism and biosynthesis, and (4) cellular carbon budgets (Hayes 1993). Autotrophic organisms assimilate inorganic carbon (marine carbonate or bicarbonate, atmospheric carbon dioxide) with d13C values typically between +2 and –8‰ while heterotrophic organisms assimilate carbon from biogenic organic matter with d13C from –5 to –35‰. However, methane with d13C values in the range of  –35 to –60‰ (thermogenic) and  –55 –85‰ (biogenic) has to be considered as a significant and sometimes unusually light carbon source for methanotrophic organisms (> Fig. 1). Typically, biomass is depleted in 13C relative to the respective carbon source due to isotope fractionation processes whose magnitudes depend on the mechanism of carbon fixation. As a consequence of differences in the isotope fractionation associated with specific biosynthetic pathways, 12C and 13C are not equally distributed between the different carbon pools (e.g., carbohydrates, proteins, lipids) of a single organism. Furthermore, it has to be taken into account that sedimentary organic matter typically represents a more or less complex mixture of biomass derived from different source organisms which all contribute their own specific carbon isotopic signature. This is revealed by compound-specific isotope analysis of biomarkers which can be attributed to specific source organisms. In general, d13C values of sedimentary organic matter are relatively similar to those of the biomass exported from the water column, indicating that even significant changes in organic matter composition during transport through the water column and early diagenesis after deposition in the sediment (including complete recycling of about 99% of the primary

Stable Isotopes in Understanding Origin and Degradation Processes of Petroleum

5

. Figure 1 Variations in stable carbon isotope ratios (VPDB standard) for different organic and inorganic natural materials (modified from Mook 2000; Schidlowski and Aharon 1992).

production) do not strongly influence the carbon isotopic composition (for review see Galimov 2006). Likewise, only small shifts towards heavier d13C values (up to 1‰) are observed for kerogens due to thermal maturation during catagenesis as a result of the release of isotopically light oil and gas (Clayton 1991). Kinetic isotope effects related to oil-to-gas cracking may lead to an enrichment of the remaining oil in 13C, which, however, is unlikely to be greater than about 4‰ (Clayton 1991). A maturation effect is also seen in the carbon isotopic composition of individual oil constituents, however, it normally will not exceed a 2–3‰ shift towards less negative values (Clayton and Bjorøy 1994). The carbon isotopic composition of bulk crude oils typically ranges between  –24 and –34‰ (> Fig. 1). Small differences in the average isotopic compositions of the C15+ fractions of saturated and aromatic hydrocarbon fractions of oils sourced from terrestrial or marine organic matter are more or less insignificant (Sofer 1984). However, it is well established that different petroleum source rocks generate oils with distinct carbon isotopic signatures. This can be illustrated by a case study from the North Viking Graben, an exploration area in the Norwegian North Sea (Gormly et al., 1994). In this area two potential source rocks, both of Upper Jurassic age, the Heather and the Draupne Formations, may contribute to the reservoired crude oils. Kerogen of the Draupne Fm. (–28 to –27‰) is isotopically slightly lighter than that of the Heather Fm. (–27 to –25‰) which corresponds well to the isotopic signatures of the generated oils (–31 to –28‰ versus –28 to –25‰). These variations in isotopic composition are furthermore correlated to those of a molecular biomarker parameter (pristane/phytane ratio) indicating an influence of the source rock depositional environment on the d13C values of the kerogens. In many cases reservoired crude oils are not derived

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from a single source but rather represent mixtures whose isotopic composition depends on the isotopic composition and relative amounts of the contributing sources. In the North Viking Graben, mixed oils sourced from both the Draupne and the Heather Fm. have d13C values in the range of –28.8 to –28.0‰ (Gormly et al., 1994). The isotopic composition of individual oil constituents, for example, gasoline-range hydrocarbons, may provide useful clues as to the mixed origin of oil (Rooney et al., 1998). Ideally, the carbon isotopic composition of mixed oil can be used to quantify the contributions of the individual sources (Peters et al., 1989). The stable carbon isotopic compositions of crude oil fractions (saturated and aromatic hydrocarbon fractions) have been shown to become enriched in 13C with decreasing geologic age (Andrusevich et al., 1998). The authors compared evolutionary changes in the biosphere to episodic changes in stable carbon isotopic compositions throughout the Phanerozoic and concluded that these isotopic shifts may be related to the diversity of preserved phytoplankton. Furthermore, crude oils generated from source rocks of Upper Jurassic age become increasingly enriched in 13C from high to low paleolatitudes (=latitudes at which the source rocks were deposited), indicating that the d13C values of oils reflect that of the primary marine biomass, which varied as a function of spatial paleoenvironmental parameters, in particular sea-surface paleotemperature (Andrusevich et al., 2000). Stable carbon isotope ratios are extremely useful for the assessment of the origin of natural gas (for review see Whiticar 1994). This is because fractionations due to kinetic isotope effects lead to a much higher variability in the d13C values of small molecules with only one to five carbon atoms, as is the case in the natural gas hydrocarbons methane (CH4), ethane (C2H6), propane (C3H8), i-butane (C4H10), n-butane (C4H10), i-pentane (C5H12) and n-pentane (C5H12). Methane is by far the predominant constituent of natural gas. The C2–C5 hydrocarbons occur in highly variable amounts which in general are low in biogenic gas and higher in thermogenic gas. The relative amount of the C2–C5 hydrocarbons in natural gas is typically referred to as the ‘‘gas wetness’’ (4). gas wetness ¼

½C2  C5  : ½C1  C5 

ð4Þ

Likewise, carbon isotopic signatures of biogenic methane are significantly more depleted in 13C than they are in thermogenic methane. Typically, d13C values of methane lighter than –60‰ and heavier than –55‰ are attributed to a purely biogenic or a purely thermogenic origin, respectively, while a mixed source has to be considered for intermediate d13C values. Methane isotopic composition provides clear evidence for the occurrence of biogenic methane in very deep reservoirs (at least down to 3,000 m) and thus is a very important hint to the existence of a subterraneous so-called deep biosphere (Schoell 1980). Recently, the isotopic compositions of ethane and propane in cold, deeply buried sediments from the southeastern Pacific were interpreted to reflect the microbial production of these hydrocarbons in situ (Hinrichs et al., 2006). The hydrogen isotopic compositions of organic components are used to a much lesser extent in petroleum geochemistry. It might be expected that they show similar fractionation behavior during processes involved in the formation and destruction of petroleum as discussed here for the distribution of the carbon isotopes. However, recent progress on this topic appears to indicate significant differences, likely due to different modes in which hydrogen is involved in biogeochemical processes. A crucial aspect seems to be that hydrogen atoms in hydrocarbons (and organic matter in general) are exchangeable with external hydrogen, both organically and inorganically bound, which likely is negligible for carbon atoms. In particular, water has to be

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considered as a relevant source of hydrogen in petroleum systems, and hydrocarbons may interact with water on the migration pathway from the source to the trap or with the formation water in a reservoir. It is true that the C–H acidity of hydrocarbons is extremely low; however, on geological timescales, i.e., many million years or even more, and at the elevated temperatures occurring in many petroleum systems, exchange reactions may become significant, although they would not be observable on a human timescale. Numerous recent studies have investigated such exchange processes and have clearly indicated that the hydrogen isotopic signatures of geological organic matter (kerogen, bulk oils, oil fractions, individual oil and gas constituents) become systematically enriched in deuterium with increasing levels of thermal maturity (Dawson et al., 2005; Lis et al., 2006; Mastalerz and Schimmelmann 2002; Pedentchouk et al., 2006; Radke et al., 2005; Schimmelmann et al., 2001; Sessions et al., 2004; Tang et al., 2005). This seems to indicate that hydrogen isotope ratios could be efficient tools in thermal history assessment where carbon isotope ratios are not very useful (see above). Beyond this, the systematics of hydrogen isotopes in organic components of petroleum systems awaits further investigations to fully establish their potential as a tool in exploration and production of petroleum.

3.2

Alteration Processes in Petroleum Reservoirs

Biodegradation in petroleum reservoirs will take place near the oil-water interface and was described to be of interest in reservoirs that were not exposed to temperatures >80 C (Connan 1984; Wilhelms et al., 2001). The oil-water contact provides conditions that are the most conducive to microbial activity. Diffusive transport of hydrocarbons through the oil column to the oil-water contact will provide electron donors, whereas inorganic nutrients required for microbial growth can be transported by water flow or diffusion in the water column to the oilwater contact (Head et al., 2003). Biodegradation processes in crude oils were described to lead to the quasi-sequential removal of compound groups as follows: n-alkanes > branched alkanes > alkylbenzenes > alkylnaphthalenes > alkylcyclohexanes, alkylphenanthrenes and alkyldibenzothiophenes > isoprenoids (C15+) > regular steranes > hopanes > aromatic steranes (Peters and Moldowan 1993; Wenger et al., 2002). However, recent work on molecular changes in biodegraded oils indicates that the degradation patterns of light hydrocarbons and n-alkanes differ in different petroleum systems. This suggests that microbial communities are different and therefore generate different molecular degradation patterns which have to be evaluated individually for each system (Elias et al., 2007). The molecular and isotopic composition of natural gas hydrocarbons (C1–C5) is used traditionally for gas-gas correlations, as has been successfully demonstrated by Boreham and coauthors for Australian gases (Boreham et al., 2001). However, influences of source, maturity and biodegradation processes on the molecular and isotopic composition of these compounds have to be evaluated quite carefully. The relative abundance of the wet gases (C2–C5) will decrease and their d13C will shift to more positive values with biodegradation. The isotopic composition of the original methane will be influenced by additional biogenic methane with light isotopic composition ( Fig. 2). The increase in the iC5/nC5 ratio is correlated to increasing d13C values of iC5 and nC5. It is obvious for all oil fields that the d13C values of iC5 are slightly more negative than the d13C values of nC5 and show a smaller enrichment with increase in the iC5/nC5 ratios (> Fig. 2). Evaluation of the carbon isotope and concentration data for iC5 and nC5 using the Rayleigh equation (2) indicates that the Rayleigh model can be applied here. It turns out

. Figure 2 d13C of iC5 (squares) and nC5 (circles) in oil samples from offshore Angola and offshore Norway (Gullfaks and Troll) plotted over their iC5/nC5 concentration ratio; data taken from Barman Skaare et al., 2007; Vieth and Wilkes 2006; Wilkes et al., 2008.

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that the isotope fractionation factors for iC5 are identical in Norway (Gullfaks) and Angola, and that the fractionation factors for nC5 are very similar (> Fig. 3) (Wilkes et al., 2008). This may indicate that the isotopic fractionation of individual hydrocarbons due to biodegradation is very similar in different petroleum systems, even if the molecular patterns of alteration are different. The carbon isotope and concentration data of the Troll oil samples have not been included in the Rayleigh evaluation because all oil samples have a relatively similar extent of biodegradation (Barman Skaare et al., 2007). For an unambiguous assessment of biodegradation a reliable non-biodegraded oil sample is necessary to give the initial isotopic composition and concentration of iC5 and nC5. The application of compound-specific isotope analysis to assess in-reservoir biodegradation has certain restrictions. Carbon isotope fractionation of the residual substrate occurs in the first irreversible reaction, which mechanistically takes place at (a) certain specific carbon atom(s) of the substrate in most known cases. Therefore, the overall isotope effect will become less with increasing number of carbon atoms in the molecule (Boreham et al., 1995). Dilution of the fractionation effect limits the application of the Rayleigh approach to light hydrocarbons (Morasch et al., 2004; Wilkes et al., 2008). It can be concluded from this that the compound-specific carbon isotope ratios of C15+ hydrocarbons will not be affected by biodegradation processes and can still be used as specific indicators of the origin of the oil as well as for oil-oil and oil-source rock correlations (Vieth and Wilkes 2006). Based on the assumption that only light hydrocarbons show carbon isotope fractionation related to biodegradation processes, it becomes clear that the carbon isotopic composition of

. Figure 3 Concentration and d13C of iC5 (squares) and nC5 (circles) for oil samples from Gullfaks (dark grey symbols; data taken from Vieth and Wilkes 2006) and an oil field offshore Angola (light grey symbols; data taken from Wilkes et al., 2008), plotted according to the Rayleigh equation as ln R/Ri over ln F. Reprinted from Wikes et al., 2008 with permission from Elsevier.

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whole oils and individual oil fractions (e.g., saturated and aromatic hydrocarbons) will not show significant changes due to biodegradation (Sofer 1984; Stahl 1980; Sun et al., 2005). Within bulk oils the isotopic signatures of components which show a significant fractionation will be overprinted by the isotopic signatures of non-degraded oil constituents becoming relatively enriched in such a process. The hydrocarbons that are present in different oil fractions (C15+ saturates and C11+ aromatics) are generally made up of larger compounds which are not expected to show significant fractionation due to biodegradation (Wilkes et al., 2008). For hydrogen isotope fractionation the same principal behavior would be expected. In general hydrogen isotope fractionation tends to be one order of magnitude larger than carbon isotope fractionation due to the higher relative mass difference between the two isotopes. Therefore, the dilution effect will become relevant at a higher number of hydrogen atoms being present within the substrate molecule. This is in agreement with results of aerobic degradation experiments where the hydrogen isotopic composition of long-chain alkanes (nC19 to nC27) did not change significantly (Pond et al., 2002). In contrast to laboratory experiments where hydrogen isotope fractionation is correlated to the decrease in substrate concentration (Mancini et al., 2003; Pond et al., 2002), the hydrogen isotope ratios of light hydrocarbons in crude oils may not show a clear dependence on biodegradation processes. It has been suggested that in petroleum reservoirs, besides differences in maturity and source, the effects of hydrogen exchange between oil and formation water over geologic times have also to be considered (Sessions et al., 2004). Thermochemical sulphate reduction (TSR) is the abiological reduction of sulphate by hydrocarbons in reservoirs close to anhydrite (source of sulphate) at high temperatures (range of minimum temperature between 100 and 140 C) (Machel 2001). Some types of hydrocarbons are more susceptible to TSR than others, for example, C2–C5 gases are more reactive than methane and saturated hydrocarbons are more reactive than aromatic hydrocarbons (Peters et al., 2005). This is confirmed by the observation that larger isotopic shifts (e.g., up to 22‰) occur during TSR for the branched and n-alkanes, whereas relatively smaller shifts (e.g., 3–6‰) have been found for the cyclic and monoaromatic hydrocarbons (Rooney 1995). Whiticar and Snowdon reported changes during TSR in d13C of individual hydrocarbons of the C5–C8 range by up to 10‰ (Whiticar and Snowdon 1999).

4

Research Needs

In order to strengthen the application of stable carbon and hydrogen isotopes as valuable process indicators for an improved understanding of petroleum generation, reservoir filling and secondary alteration, it is necessary to deepen the insight into the hydrogen exchange processes occurring in petroleum reservoirs over geological time scales. This will help to evaluate compound-specific hydrogen isotope ratios of petroleum hydrocarbons. Of further interest are the chances and limitations of 2-dimensional compound-specific isotope analysis (13C, D) (Fischer et al., 2008; Zwank et al., 2005) and its application in petroleum reservoir studies. Biodegradation processes can be evaluated more easily if the fundamental kinetic isotope effects of biodegradation reactions occurring during anaerobic degradation of hydrocarbons have been deciphered. This will reduce the intensive work that is necessary to obtain isotope fractionation factors that are applicable in reservoir studies.

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References Andrusevich VE, Engel MH, Zumberge JE, Brothers LA (1998) Secular, episodic changes in stable carbon isotope composition of crude oils. Chem Geol 152: 59–72. Andrusevich VE, Engel MH, Zumberge JE (2000) Effects of paleolatitude on the stable carbon isotope composition of crude oils. Geology 28: 847–850 Barman Skaare B, Wilkes H, Vieth A, Rein E, Barth T (2007) Alteration of crude oils from the Troll area by biodegradation: Analysis of oil and water samples. Org Geochem 38: 1865–1883 Boreham CJ, Dowling LM, Murray AP (1995) Biodegradation and maturity influences on n-alkane isotopic profiles in terrigenous sequences. In Organic Geochemistry: Developments and applications to energy, climate, environment and human history, Selected paper of the 7th International Meeting on Organic Geochemistry. JO Grimalt and C Dorronsoro (eds.) pp 539–544. Boreham CJ, Hope JM, Hartung KB (2001) Understanding source, distribution and preservation of Australian natural gas; a geochemical perspective. In: 2001 APPEA conference Australian Petroleum Production and Exploration Association, Canberra, Australia, pp 523–547. Clark ID, Fritz P (1997) Environmental isotopes in hydrogeology. Boca Raton, FL: Lewis Publishers. Clayton CJ (1991) Effect of maturity on carbon isotope ratios of oils and condensates. Org Geochem 17: 887–899. Clayton CJ, Bjorøy M (1994) Effect of maturity on 13 12 C/ C ratios of individual compounds in North Sea oils. Org Geochem 21: 737–750. Connan J (1984) Biodegradation of crude oils in reservoirs. In Advances in petroluem geochemistry, vol. 1. JM Brooks, D Welte (eds.). London: Academic Press, pp 299–355. Craig H (1957) Isotopic standards for carbon and oxygen and correction factors for mass-spectrometric analysis of carbon dioxide. Geochim Cosmochim Acta 12: 133–149. Dawson D, Grice K, Alexander R (2005) Effect of maturation on the indigenous dD signatures of individual hydrocarbons in sediments and crude oils from the Perth Basin (Western Australia). Org Geochem 36: 95–104. Elias R, Vieth A, Riva A, Horsfield B, Wilkes H (2007) Improved assessment of biodegradation extent and prediction of petroleum quality. Org Geochem 38: 2111–2130. Fischer A, Herklotz I, Herrmann S, Thullner M, Weelink SAB, Stams AJM, Schlo¨mann M,

Richnow HH, Vogt C (2008) Combined carbon and hydrogen isotope fractionation investigations for elucidating benzene biodegradation pathways. Environ Sci Technol 42: 4356–4363. Galimov EM (2006) Isotope organic geochemistry. Org Geochem 37: 1200–1262. George SC, Boreham CJ, Minifie SA, Teerman SC (2002) The effect of minor to moderate biodegradation on C5 to C9 hydrocarbons in crude oils. Org Geochem 33: 1293–1317. Gormly JR, Buck SP, Chung HM (1994) Oil-source rock correlation in the North Viking Graben. Org Geochem 22: 403–413. Hagemann R, Nief G, Roth E. (1970) Absolute isotopic scale for deuterium analysis of natural waters. Absolute D/H ratio for SMOW. Tellus 22: 712–715. Hayes JM (1993) Factors controlling 13C contents of sedimentary organic compounds: Principles and evidence. Mar Geol 113: 111–125. Head IM, Jones DM, Larter SR (2003) Biological activity in the deep subsurface and the origin of heavy oil. Nature 426: 344–352. Hinrichs KU, Hayes JM, Bach W, Spivack AJ, Hmelo LR, Holm NG, Johnson CG, Sylva SP (2006) Biological formation of ethane and propane in the deep marine subsurface. Proc Natl Acad Sci USA 103: 14684–14689. Hoefs J (2004) Stable isotope geochemistry. Berlin: Springer. Kniemeyer O, Musat F, Sievert SM, Knittel K, Wilkes H, Blumenberg M, Michaelis W, Classen A, Bolm C, Joye SB, Widdel F (2007) Anaerobic oxidation of short-chain hydrocarbons by marine sulphatereducing bacteria. Nature 449: 898–902. Lis GP, Schimmelmann A, Mastalerz M (2006) D/H ratios and hydrogen exchangeability of type-II kerogens with increasing thermal maturity. Org Geochem 37: 342–353. Machel HG (2001) Bacterial and thermochemical sulfate reduction in diagenetic settings – old and new insights. Sediment Geol 140: 143–175. Mancini SA, Ulrich AC, Lacrampe-Couloume G, Sleep B, Edwards EA, Sherwood Lollar B (2003) Carbon and hydrogen isotopic fractionation during anaerobic biodegradation of benzene. Appl Environ Microbiol 69: 191–198. Mastalerz M, Schimmelmann A (2002) Isotopically exchangeable organic hydrogen in coal relates to thermal maturity and maceral composition. Org Geochem 33: 921–931. Masterson WD, Dzou LIP, Holba AG, Fincannon AL, Ellis L (2001) Evidence for biodegradation and evaporative fractionation in West Sak, Kuparuk

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and Prudhoe Bay field areas, North Slope, Alaska. Org Geochem 32: 411–441. McKinney CR, McCrea JM, Epstein S, Allen HA, Urey HC (1950) Improvements in mass spectrometers for the measurement of small differences in isotope abundance ratios. Rev Sci Instrum 21: 724–730. Milkov AV, Dzou L (2007) Geochemical evidence of secondary microbial methane from very slight biodegradation of undersaturated oils in a deep hot reservoir. Geology 35: 455–458. Mook WG (2000) Environmental isotopes in the hydrological cycle – Principles and applications. Paris: UNESCO. Morasch B, Richnow HH, Vieth A, Schink B, Meckenstock RU (2004) Stable isotope fractionation caused by glycyl radical enzymes during bacterial degradation of aromatic compounds. Appl Environ Microbiol 70: 2935–2940. Pallasser RJ (2000) Recognizing biodegradation in gas/oil accumulations through the d13C compositions of gas components. Org Geochem 31: 1363–1373. Pedentchouk N, Freeman KH, Harris NB (2006) Different response of dD values of n-alkanes, isoprenoids, and kerogen during thermal maturation. Geochim Cosmochim Acta 70: 2063–2072. Peters KE, Moldowan JM, Driscole AR, Demaison GJ (1989) Origin of Beatrice oil by co-sourcing from Devonian and Middle Jurassic source rocks, inner Moray Firth, United Kingdom. AAPG Bull 73: 454–471. Peters KE, Moldowan JM (1993) The biomarker guide: interpreting molecular fossils in petroleum and ancient sediments. New Jersey: Prentice Hall. Peters KE, Walters CC, Moldowan JM (2005) The biomarker guide – Biomarkers and isotopes in the environment and human history. Cambridge, UK: Cambridge University Press. Pond KL, Huang Y, Wang Y, Kulpa CF (2002) Hydrogen isotopic composition of individual n-alkanes as an intrinsic tracer for bioremediation and source identification of petroleum contamination. Environ Sci Technol 36: 724–8. Radke J, Bechtel A, Gaupp R, Puttmann W, Schwark L, Sachse D, Gleixner G (2005) Correlation between hydrogen isotope ratios of lipid biomarkers and sediment maturity. Geochim Cosmochim Acta 69: 5517–5530. Rooney MA (1995) Carbon isotope ratios of light hydrocarbons as indicators of thermochemical sulfate reduction. In Organic Geochemistry: Developments and applications to energy, climate, environment and human history, JO Grimalt and C Dorronsoro (eds.) Selected paper of the 7th International Meeting on Organic Geochemistry, pp 523–525.

Rooney MA, Vuletich AK, Griffith CE (1998) Compoundspecific isotope analysis as a tool for characterizing mixed oils: An example from the West of Shetlands area. Org Geochem 29: 241–254. Schidlowski M, Aharon P (1992) Carbon cycle and carbon isotope record: geochemical impact of life over 3.8 Ga of earth history. In: Early organic evolution: Implications for mineral and energy resources. M Schidlowski (ed.). Berlin: Springer, pp 147–175. Schimmelmann A, Boudou JP, Lewan MD, Wintsch RP (2001) Experimental controls on D/H and 13C/12C ratios of kerogen, bitumen and oil during hydrous pyrolysis. Org Geochem 32: 1009–1018. Schoell M (1980) The hydrogen and carbon isotopic composition of methane from natural gases of various origins. Geochim Cosmochim Acta 44: 649–661. Sessions AL, Sylva SP, Summons RE, Hayes JM (2004) Isotopic exchange of carbon-bound hydrogen over geologic timescales. Geochim Cosmochim Acta 68: 1545–1559. Sharp Z (2006) Principles of Stable Isotope Geochemistry Upper Saddle River, NJ: Prentice Hall. Sofer Z (1984) Stable carbon isotope compositions of crude oils; application to source depositional environments and petroleum alteration. AAPG Bull 68: 31–49. Stahl WJ (1980) Compositional changes and 13C/12C fractionations during the degradation of hydrocarbons by bacteria. Geochim Cosmochim Acta 44: 1903–1907. Sun Y, Chen Z, Xu S, Cai P (2005) Stable carbon and hydrogen isotopic fractionation of individual n-alkanes accompanying biodegradation: evidence from a group of progressively biodegraded oils. Org Geochem 36: 225–238. Tang Y, Huang Y, Ellis GS, Wang Y, Kralert PG, Gillaizeau B, Ma Q, Hwang R (2005) A kinetic model for thermally induced hydrogen and carbon isotope fractionation of individual n-alkanes in crude oil. Geochim Cosmochim Acta 69: 4505–4520. Vieth A, Wilkes H (2006) Deciphering biodegradation effects on light hydrocarbons in crude oils using their stable carbon isotopic composition: a case study from the Gullfaks oil field, offshore Norway. Geochim Cosmochim Acta 70: 651–665. Welte DH, Kratochvil H, Rullko¨tter J, Ladwein H, Schaefer RG (1982) Organic geochemistry of crude oils from the Vienna Basin and an assessment of their origin. Chem Geol 35: 33–68. Wenger LM, Davis CL, Isaksen GH (2002) Multiple controls on petroleum biodegradation and impact on oil quality. SPE Reservoir Eval Eng 5: 375–383. Whiticar MJ (1994) Correlation of natural gases with their sources. In: The petroleum system – from source to trap American Association of Petroleum

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Wilkes H, Vieth A, Elias R (2008) Constraints on the quantitative assessment of in-reservoir biodegradation using compound specific stable carbon isotopes. Org Geochem 39: 1215–1221. Zwank L, Berg M, Elsner M, Schmidt TC, Schwarzenbach RP, Haderlein SB (2005) New evaluation scheme for two-dimensional isotope analysis to decipher biodegradation processes: application to groundwater contamination by MTBE. Environ Sci Technol 39: 1018–1029.

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6 The Microbial Production of Methane and Other Volatile Hydrocarbons M. Formolo Max Planck Institute for Marine Microbiology, Bremen, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 114

2 Microbial Formation of Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 2.1 Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115 2.2 Syntrophic Communities and Methane Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 119 3 Microbial Formation of Non-Methane Volatile Hydrocarbons . . . . . . . . . . . . . . . . . . . 120 3.1 Non-Methane Short Chain Volatile Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 121 4

Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122

5

Research Needs and Critical Knowledge Gaps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_6, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: In the mid-1930s, scientists were attempting to unravel the mechanisms responsible for microbial methane production. Following in the mid-1950s, scientists began assigning a microbial origin to volatile hydrocarbon gas other than methane. Since these earliest reports, biogenic methane and other volatile hydrocarbons have been measured in terrestrial, marine and lacustrine environments. Decades of research have demonstrated that these processes are complex, involving multiple organisms and a plethora of enzymatic processes. However, when combined in the appropriate context, these microbial processes highlight the exciting, important, and unique biogeochemical niche these organisms fill. This review attempts to integrate the microbial and geochemical conditions promoting microbial production of volatile hydrocarbons. As continued research focuses on understanding the environmental and biological limits of biogenic gas production, it appears that these communities have the ability to adapt and survive in the most energy-limited conditions on Earth. The dominant regions of microbial volatile hydrocarbon production are usually restricted to zones with specific geochemical conditions. Biogenic methane production is almost exclusively found in environments where sulfate is depleted and adequate substrates for methanogenesis are available. Conditions leading to the formation of other short-chain volatile hydrocarbons are less well understood. Biogenic methane production requires a hierarchal transfer of energy and substrates among a variety of organisms culminating in terminal carbon remineralization by methanogenic archaea. Recently, the recognition of other microbially produced non-methane hydrocarbons - specifically ethane, propane, and isoprene -has demonstrated the importance of these compounds in many environments. While the mechanisms for the formation of these non-CH4 hydrocarbons are not as well understood as their C1 counterpart, there are likely similarities in the metabolic and geochemical conditions. At the conclusion of this minireview, a series of basic questions and needs regarding future research objectives regarding the microbial production of methane and non-methane hydrocarbons are proposed, including developing a more comprehensive understanding of the mechanisms leading to their formation and degradation.

1

Introduction

Volatile organic compounds have a high enough vapor pressure to vaporize at standard temperature and pressure conditions. These volatile compounds include C1-C7 saturated hydrocarbons, alkenes, aromatics, and other non-hydrocarbon volatile organic compounds. Since several of these compounds are found within marine and terrestrial sediments and the atmosphere, their formation and distribution are important to many Earth surface processes, including climate change, extinction events, petroleum and gas deposits, and atmospheric pollution. Of particular interest are the microbially produced volatile hydrocarbons. The most common microbially produced gas is methane, which accounts for 85% of the annual global methane production (Valentine et al., 2004). Methane is in greater abundance in the atmosphere than non-methane volatile hydrocarbons due to the relative stability of CH4 versus other short-chain hydrocarbons such as ethane and propane. For example, the reactivity of non-methane hydrocarbons is approximately two orders of magnitude greater than for methane, resulting in rapid turnover and oxidation in the atmosphere. Calculated residence times for ethane range from 1 to 2 months, while propane and high molecular weight volatile hydrocarbons have a residence time of less than 1 day, compared with the residence time of methane which is approximately 8–12 years (See > Chapter 18, Vol. 1, Part 4). These short

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6

residence times prevent build-up of these non-methane hydrocarbons in the atmosphere. Though short-lived, these volatile hydrocarbons have a large impact on Earth system processes (Bonsang and Lambert, 1985; Whiticar, 1990). In addition to their atmospheric residence time, these volatile hydrocarbons are also susceptible to rapid oxidation in aqueous environments, either biologically or abiotically (Rudd et al., 1974), under aerobic or anaerobic conditions. While many mechanisms can produce and consume these compounds, only the microbial processes involved with the production and consumption of permanently gaseous hydrocarbons, specifically methane, and non-CH4 volatile hydrocarbons will be considered in this mini-review.

2

Microbial Formation of Methane

2.1

Methane

Previous discussions and reviews (see Schoell, 1988) pertaining to the origin of hydrocarbon gases have drawn a distinction between biogenic and microbially produced volatile hydrocarbons. Throughout this short review, all biogenic hydrocarbons will be assumed to be the result of microbial processes. Microbial sources of methane had been recognized prior to other non-methane hydrocarbons (Barker, 1936). The recognition of biogenic gas production other than methane was only made recently during the middle of the 20th century (Davis and Squires, 1954). Since these initial reports, the recognition of biogenic gas production and consumption has continued to expand, highlighting the importance of such processes in almost all environments. The formation of biogenic gas is often the direct result of the biodegradation of organic matter and can occur under anaerobic and aerobic conditions in terrestrial, lacustrine, and marine environments. These environments include swamps, rice paddy fields, anoxic freshwater lakes, sublittoral-marine sediments, glacial drifts, marine sediments (shallow and deep), deep sedimentary basins, salt marshes, coal deposits, petroleum reservoirs, peat bogs, and even the surface ocean (Barnes and Goldberg, 1976; Claypool and Kvenvolden, 1983; Hinrichs et al., 2006; Martens and Berner, 1974; Oremland et al., 1982a; Rashid and Vilks, 1977; Shoell, 1980, 1988; Whiticar, 1990, Karl et al., 2008). The dominant hydrocarbon gas that is produced in many of these environments is methane, with minor amounts of other gases. Some biogenic gases are also produced during critical metabolic processes, e.g. isoprene, and will be discussed later. All known methanogens are obligately anaerobic archaea. Growth substrates for methanogenic archaea include a variety of compounds including CO2 + H2, formate, acetate, methanol, ethanol, benzoate, carbon monoxide, ethylated sulfur compounds, and methylated amines (Barker, 1936; Hippe et al., 1979; King et al., 1983; Nottingham and Hungate, 1969; Smith and Mah, 1978; Stadtman and Barker, 1949; Winfrey and Ward, 1983; Zeikus et al., 1985). Under some environmental conditions it has been suggested that methionine, methyl mercaptan, and dimethyl sulfides may be alternative carbon substrates for methane production (Oremland et al., 1982b; Zinder and Brock, 1978). While a variety of compounds may be utilized as substrates during methanogenesis, the majority of natural environments do not contain sufficient concentrations of these compounds to maintain methanogenesis. Globally, acetate and CO2 + H2 are the dominant substrates fueling methanogenesis in the majority of natural systems. The formation of biogenic gas dominantly occurs during the early stages of sediment burial and at low temperatures ( Fig. 1) (Conrad, 1989; Games and Hayes, 1978; Jenden and Kaplan, 1986; Schoell et al., 1983; Whiticar, 1983). Simplified reactions of these pathways are: CH3 COO þ H2 O ! CH4 þ HCO 3

ð1Þ

þ HCO 3 þ 4H2 þ H ! CH4 þ 3H2 O

ð2Þ

Both these mechanisms occur in marine and lacustrine sediments, though CO2-reduction dominates in marine sediments and aceticlastic methanogenesis in freshwater sediments (Claypool and Kaplan, 1974; Whiticar et al., 1986). During acetate fermentation, both the CH4 and the CO2 are primarily derived from the methyl group of acetate. Two different processes can result in the consumption of acetate and the formation of methane; these are aceticlastic methanogenesis and a syntrophic relationship between acetate oxidation and hydrogenotrophic methanogenesis (Barker, 1936; Hattori, 2008; Schink, 1997; Zinder and Koch, 1984). The methane formed during CO2-reduction is derived from the in-situ pore-water bicarbonate, or carbonate alkalinity. These substrates are often remnants of earlier carbon remineralization processes such as microbial organoclastic and methanotrophic sulfate reduction. The terminal enzymatic processes during both aceticlastic and hydrogenotrophic methanogenesis involve the key enzyme Methyl-coenzyme M reductase. However, during CO2reduction, the initial activation and transfer of the C1 unit from the substrate involves the coenzyme methanofuran. During aceticlastic methanogenesis, the formation of CH4 from acetate is initiated by acetyl-coezyme A (Ferry, 1992; Weimer and Zeikus, 1978). Following this activation, further enzymatic processes create a methyl-coenzyme M complex that can be further reduced to methane. In addition, transition metals such as nickel and cobalt are important for the enzymatic processes involved in methanogenesis (Hausrath et al., 2007; Kida et al., 2001; Scho¨nheit et al., 1979). Specifically, nickel is the core transition metal in coenzyme F430, which is an essential component of the methyl reductase structure (Ermler et al., 1997). Acetate fermentation may be the dominant contributor to the formation of methane; however, the process is considerably slower than the formation of methane via CO2-reduction

The Microbial Production of Methane and Other Volatile Hydrocarbons

6

. Figure 1 Anaerobic degradation of organic matter and the formation of biogenic methane.

(Blaut, 1994). These two processes – acetate fermentation and CO2-reduction – yield free energy and are exogenic, and the free energy gain can be used to synthesize ATP. According to 0 Oremland (1988), the formation of methane via acetate fermentation yields DGo  = 31 kJ 0 1 1 mol and CO2-reduction yields DGo  = 135 kJ mol . The energy yield for methane formation from CO2 reduction is approximately 4.3 times greater. This gain in free energy during CO2-reduction is sufficient for synthesis of at least one molecule of ATP. This overall energy gain, or energy conservation, during methanogenesis is due to the enzymatic activity of the terminal process during methanogenesis involving methyl reductase enzyme. The activity of these enzymatic processes is critical to the maintenance and preservation of methanogenic communities, and allows for energetically favorable metabolisms under energetically deficient environments. The relative contribution of hydrogenotrophic or aceticlastic methanogenesis to methane formation can vary depending on environmental conditions. Ferry (1992) and Valentine et al. (2004) suggested that 70% of the global anaerobic biogenic methane formation is driven by acetate, though CO2-reduction dominates in shallow marine sediments. Valentine et al. (2004) and Whiticar (1999 and references therein) determined that in terrestrial and lacustrine

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environments, aceticlastic methanogenesis accounts for 70% of all methane produced. Incubation studies by Fey et al. (2003) demonstrated that at moderate temperatures, e.g. 50 C, the methane production is dominated by CO2-reduction. They also showed a transition between the two dominant methanogenic pathways related to variations in temperature. Recent research in terrestrial ecosystems, such as rice paddies, indicate as much as 75–80% of the methane emitted is derived from aceticlastic methanogenesis (Roy et al., 1997). The active metabolic mechanism is controlled by the presence or absence of sulfate in the sediments. During microbial sulfate reduction, which occurs dominantly in marine environments, the compounds acetate, formate, and occasionally CO2 + H2 are competitive aceticlastic substrates and are preferentially utilized by sulfate reducers, eliminating the opportunity for aceticlastic methanogens to survive (Martens and Berner, 1974; Whiticar et al., 1986; Whiticar, 1999). In marine and estuarine sediments where sulfate concentrations are greater than 200 mM, methanogenesis is inhibited (Martens and Berner, 1974; Oremland and Polcin, 1982c). While the presence of sulfate and sulfate reducing organisms inhibits methanogenic activity, the products of sulfate reduction can lead to the proper chemical conditions for methanogens. During sulfate reduction, the continued remineralization of organic matter leads to increased dissolved inorganic carbon, or bicarbonate pools. Following the depletion of sulfate and the competitive substrate acetate, the elevated bicarbonate concentrations provide the necessary substrate for the biogenic methane production via CO2-reduction (Whiticar et al., 1986; Whiticar, 1999). While in freshwater systems, which are characterized by low sulfate concentrations, acetate fermentation may dominate due to the absence of competitive microbial communities (Winfrey and Zeikus, 1977). The absence of sulfate reducers allows for the buildup of short-chain volatile fatty acids, specifically acetate, which provide the appropriate substrates for methanogenesis (Whiticar et al., 1986; Whiticar, 1999). Also, the presence of methyl amines in these lacustrine sediments may fuel methanogenesis. In anoxic lacustrine sediments where aceticlastic methanogenesis may deplete the acetate pool, CO2-reduction can also occur- though its relative contribution is minor. In environments other than marine and lacustrine sediments, microbial methane production can occur via non-competitive substrate mechanisms. These mechanisms also include acetate-utilizing and methylotrophic pathways. However, the roles and global contributions of methane from these mechanisms are not well constrained. One environment that is of particular interest are oil reservoirs (See > Chapter 54, Vol. 4, Part 6). Field-evidence indicates CO2-reduction is the dominant methanogenic process in petroleum reservoirs, however, recent models based on thermodynamic considerations suggest that the oxidation of alkanes to acetate alone is linked to acetoclastic methanogenesis in these environments (Dolfing et al., 2008). Terrestrial environments including rice paddies, salt marshes, and wetlands harbor microbial communities that are capable of these metabolisms. Many of these environments are extremely sensitive to climate perturbations. These environments are often implicated as major sources of methane to the atmosphere during climate change throughout Earth history (Andreae and Schimel, 1989; Cicerone and Oremland, 1988; Whiticar, 1990, 1999). In addition to the presence of competitive and non-competitive substrates influencing the mechanism of microbial methanogenesis, the nature of organic matter may also affect the metabolic pathway. Conrad (1999, 2005) demonstrated that organic matter variations can vary the relative contributions of methane from aceticlastic methanogenesis and CO2-reduction. According to Conrad (1999, 2005), if polysaccharides are degraded, the theoretical contribution of methane production acetate fermentation would be approximately 70%. The input of organic matter into lacustrine, wetlands, rice paddies, and marine

The Microbial Production of Methane and Other Volatile Hydrocarbons

6

environments can be vastly different, and these variations may be an important mechanism in controlling the process of methane production. Additionally, the initial degradation may be critical in determining the methanogenic pathway. Conrad (2005) also suggests that if homoacetogenesis is the dominant or initial fermentation process degrading organic matter, then it would be possible to have an even higher contribution from acetate fermentation. While the examples of these controls are scarce, it is apparent that the type and initial degradation processes can be important regulators influencing the dominant methanogenic process. Temperature, the availability of substrates and nutrients, primary degradation processes, and organic matter composition determine the dominant methane producing mechanism. It also appears that the age of the organic matter may play an important role. Chanton et al. (1995) proposed that the relative age of organic matter in different environments may determine whether acetate fermentation or CO2-reduction is dominant. Chanton et al. (1995) and Nakagawa et al. (2002) concluded that in relatively fresh and young 14C-enriched organic matter the dominant methanogenic mechanism is aceticlastic methanogenesis. The authors also concluded that in older, 14C-depleted, organic matter dominated environments, CO2-reduction is the dominant mechanism. The younger, fresher organic matter may contain a higher abundance of acetyl moieties that may be liberated by hydrolysis or homoacetogenic fermentation that stimulates and favors acetate fueled methane production. The 14C-depleted, or aged, organic matter has likely been subjected to both biological attack and diagenetic alteration, leaving it deficient in acetyl moieties.

2.2

Syntrophic Communities and Methane Formation

The metabolic energy available during methanogenesis is minimal. As a result of the minimal energy gains often encountered during methanogenesis, it can be energetically favorable for methanogens to participate in mutually beneficial syntrophic relationships with other organisms. A mere 15% of the total energy available during aerobic oxidation is released during the conversion of a simple monosaccharide (hexose) to methane and carbon dioxide during anaerobic degradation. These low energy yields require unique adaptations among microbial communities. In addition to these minimal free energy gains, only a small percentage of the available organic matter or substrates are actually accessible to methanogens (e.g. CO2 and acetate). This restricted access to energy has led to the evolution of syntrophic partnerships between methanogens and other organisms that can access the available substrates. Overall, the production of methane is the culmination of a hierarchal syntrophic degradation of organic matter (See > Chapter 22, Vol. 1, Part 5). According to Schink (1997) and Hattori (2008), under conditions of light or oxidant (e.g. oxygen, ferric iron, or sulfate) deficiency it would require at least four physiologically different microbial groups to achieve organic matter degradation leading to biogenic methane formation (> Fig. 1). Compounds which may be accessible under such syntrophic conditions include high-molecular weight organic compounds such as polaysachharides, glucose, proteins, fats or long-chain saturated hydrocarbons, and aromatic compounds. The following is an example of how the compounds can be degraded, or shuttled through this syntrophic hierarchy. The first step of the relationship requires the degradation of complex organic matter such as cellulose by hydrolytic or cellulolytic bacteria during hydrolysis. This hydrolysis creates simple monomers, such as sugars, amino acids, and long-chain fatty acids. These byproducts of hydrolysis are then fermented by the primary fermenting bacteria into alcohols, short-chain fatty acids, acetate, CO2, and H2.

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Another group of organisms-secondary fermenting bacteria-can subsequently convert, via oxidation, the reduced substrates to acetate and H2 + CO2. These compounds are more readily available to both acetoclastic and hydrogenotrophic methanogens. The continual removal of the produced hydrogen and the subsequent low hydrogen partial pressures from these secondary fermenting organisms by methanogens is a critical process resulting in a beneficial and efficient syntrophy among these various groups of organisms. This syntrophy results in the efficient transfer and gain of metabolic energy among the microbial communities that ultimately culminates in the terminal remineralization of organic matter, influencing both modern and ancient global biogeochemical cycles.

3

Microbial Formation of Non-Methane Volatile Hydrocarbons

Methane is the dominant microbially produced volatile hydrocarbon on Earth; however, other non-methane volatile hydrocarbons are important components of many Earth system processes. Examples of non-CH4 microbially produced volatile hydrocarbons include ethane, propane, butane, ethylene, ethene, propene, and isoprene (See > Chapter 7, Vol. 1, Part 2). Recognizing and assigning a microbial origin to these non-methane volatile hydrocarbons has been a difficult challenge. For example, it was commonly believed that volatile short chain hydrocarbon gases other than methane were derived solely from a thermogenic source, particularly in deep marine sediments and terrestrial gas reservoirs. This observation led to the development and utilization of diagnostic geochemical proxies, such as gas wetness, as an indicator of gas source (Bernard et al., 1978). While the conclusions drawn from these indicators are often correct the continued refinement and development of diagnostic geochemical signatures, including stable isotope signatures, has demonstrated that gases traditionally assigned a thermogenic origin may indeed be biogenic - specifically microbial. Hinrichs et al. (2006) analyzed deep marine sediments from Leg 201 of the Ocean Drilling Program and concluded that ethane and propane measured in the sediments were microbial in origin. Prior to this recent publication, results from decades of ocean drilling expeditions suggested that 99% of all gas detected in deeper oceanic sediment was methane (Claypool and Kvendvolden, 1983). Other observations of non-methane short chain volatile hydrocarbons were reluctantly assigned a thermogenic origin, including earlier reports from the coastal sediments of Peru (Whiticar and Suess, 1990) and Spitsbergen (Knies et al., 2004). However, these gases may indeed be microbial in origin. While non-CH4 volatile hydrocarbon gases are scarce in the deep ocean, there were many reports suggesting the microbial origins of non-methane volatile hydrocarbons in shallow marine, terrestrial, and anoxic estuarine sediments. As early as 1954, Davis and Squires suggested that C2, C2:1, C3, and C3:1 gases can be of microbial origin (Davis and Squires, 1954). Following this report, Emery and Hoggan (1958) measured C2 through C7 hydrocarbons in sediments from the Santa Barbara Basin and concluded that these gases were result of microbial processes. Following these seminal reports of biogenic non-methane gas, Hunt (1974) demonstrated that marine organisms from the Black Sea do indeed produce ethene and propene. Early incubation studies of anoxic estuarine sediments led to the production of ethene, propane, isobutene, and butane (Vogel et al., 1982), and these incubations suggested a biological source of gas. The mounting evidence from these various observations led to the suggestion that biogenic production of these gases was a likely and viable source of these hydrocarbons (Claypool and Kvenvolden, 1983; Kvenvolden and Redden, 1980; Whelan et al., 1980).

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6

Initially; it could not be confirmed that these observations were indeed due to the microbial production of these gases, though it was suggested (Bernard et al., 1978; Emery and Hoggan, 1958). It was not until Whelan et al. (1980) and Kvenvolden and Redden (1980) that the origins of many of these gases in anoxic sediments were proven to be microbial. While there was mounting evidence of a microbial origin for many non-CH4 volatile hydrocarbons, the mechanisms of their production were, and still remain, poorly understood. Since the initial reports focusing on anoxic sediments, the microbial origin of gases has been recognized in a wide range of other environments, including deep marine sediments, deep terrestrial basins, soil sediments, and most recently the aerobic photic zone water column (Karl et al., 2008). In these environments, the most dominant non-methane volatile hydrocarbons that are produced are ethane, ethylene, propane, propene, butane, butene, and isoprene (for a complete review, refer to Ladygina et al., 2006 and references therein). These compounds are generated by a wide range of microorganisms including aerobic and anaerobic bacteria, archaea, cyanobacteria, and yeasts. Organisms produce these compounds for both ecological and physiological functions. In many cases, the specific reasons why these compounds are formed and their specific utility remain elusive; however, evidence suggests that these components are integral in regulating growth, repelling predators, and signaling molecules during intra- or interspecies interactions (Ladygina et al., 2006).

3.1

Non-Methane Short Chain Volatile Hydrocarbons

Microbial ethane and ethylene production have been documented in a variety of soil environments (Fukuda et al., 1984; Smith and Cook, 1974). It has even been suggested that in anoxic estuarine sediments, ethane may be formed by methanogens (Oremland, 1981). Propane, propylene, butane, isobutene, 1-butene, trans-2-butene, and cis-2-butene were recognized as microbial metabolites under aerobic conditions (Fukuda et al., 1984); and recently, Hinrichs et al. (2006) demonstrated that ethane and propane may be microbially produced in deep oceanic sediments. The biosynthesis of these volatile non-CH4 hydrocarbons still constitutes an important area of research. A recent review by Ladygina et al. (2006) summarizes the most current knowledge regarding the microbial biosynthesis of many hydrocarbons, including volatile non-methane hydrocarbons. Three of the most well-understood mechanisms are the formation of ethylene, isoprenoids, and isobutene. The biosynthesis of ethylene can occur via three different pathways. The most common pathway is referred to as the KMBA pathway and requires the transformation of the a-amino acid methionine to 2-keto-4-methylthiobutyric acid (KMBA). In the presence of hydroxyl radicals, the KMBA is oxidized to ethylene (Ladygina et al., 2006). Ethylene is also the byproduct of the reduction of acetylene by nitrogenase. The formation of isobutene also directly involves the alteration of an amino acid, l-leucine, via a decarboxylation reaction that produces isobutene as a final product (Fukuda et al., 1985). Isoprene is one of the more common volatile non-methane microbially produced hydrocarbons, though its formation, utility, and release as a volatile organic compound are not well understood. Horbach et al. (1993) proposed the mechanisms for the biosynthesis of isoprenoids in bacteria by monitoring the formation of metabolic intermediates during isoprenoid formation. The authors concluded that there are at minimum two distinct pathways responsible for the formation of isoprenoids-these being the MVA (mevalonate) and the MEP

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(methylerythritol phosphate) pathway. There appears to be a direct connection between isoprene release and isoprenoid formation (Wagner et al., 1999). The initial report of bacterial isoprene production was published by Kuzma et al. (1995). Subsequently, others have demonstrated isoprene production from a variety of Gram-positive and Gram-negative bacteria, with Bacillus appearing to produce the highest concentrations (Kuzma et al., 1995; Wilkins, 1996). The optimum temperature for isoprene production was 45
C, leading the authors to suggest an enzymatic control (Kuzma et al., 1995). Wagner et al. (1999) attempted to determine the mechanisms responsible for the release of isoprene. Monitoring three separate phases of bacterial growth, Wagner et al. (1999) concluded that during the formation and release of isoprene, the released isoprene is a compensation mechanism for a metabolic overflow, or the isoprene is in surplus during isoprenoid formation. The authors also suggested that release of isoprene could occur when the transfer of the isoprene carbon to higher isoprenoids is restricted. This surplus or restriction could result in the bacteria releasing the isoprene, and due to the high volatility of isoprene, this would be a simple release mechanism. The bacterial formation and release of isoprene may also be significant as predator repellents, growth inhibitors and promoters, communication agents, or in other unknown capacities. For a more detailed review regarding the biosynthesis of isoprene and other non-CH4 volatile hydrocarbons, refer to Ladygina et al. (2006) (See > Chapter 12, Vol. 2, Part 3). Hinrichs et al. (2006) documents the first evidence of biogenic ethane and propane formation in the deep marine subsurface up to 380 m below the seafloor. Using stable isotope measurements and thermodynamic calculations based on in-situ methane and acetate concentrations it was demonstrated that ethanogenic and propanogenic organisms could successfully compete with acetoclastic methanogens for acetate. The proposed mechanisms for ethanogenesis (3) and propanogenesis (4) are as follows: CH3 COO þ 3H2 þ Hþ ! C2 H6 þ 2H2 O

ð3Þ

þ CH3 COO þ HCO 3 þ 6H2 þ 2H ! C3 H8 þ 5H2 O

ð4Þ

Hinrichs et al. (2006) suggested that biogenic ethane could be formed by an ethylated Coenzyme-M, an enzyme almost identical to the terminal enzyme in methanogenesis (Oremland, 1981); or, if present, ethanethiol could be a precursor (Oremland et al., 1988). For propanogenesis, Hinrichs et al. (2006) documents that propanethiol, a common substrate for biogenic propane production (Oremland et al., 1988), was not measured and no other mechanism of formation is currently understood. These new findings highlight the importance, and likely ubiquitous production, of microbially produced volatile non-methane hydrocarbons in the majority of Earth’s environments.

4

Summary

Since the early recognition of microbially produced volatile hydrocarbons, significant advances have elucidated many of the complex enzymatic processes and environmental conditions that promote their production. In addition to microbial CH4 production it is apparent that there is a wide range of important and volatile non-CH4 hydrocarbons that are microbially produced. While the majority of previous research has focused on microbial methane, likely the result of economic and climate focused studies, new and valuable insights and discoveries are highlighting the important mechanisms and processes responsible for the microbial

The Microbial Production of Methane and Other Volatile Hydrocarbons

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production of these other non-CH4 hydrocarbons. Further research will continue to discern the mechanisms responsible for this biogenic gas and the critical role it plays in past, present, and future Earth processes.

5

Research Needs and Critical Knowledge Gaps

The microbial production of volatile hydrocarbons is an important biogeochemical process. While a vast reservoir of knowledge regarding the biogenic formation and consumption of methane and non-methane hydrocarbons exists and is expanding with new and exciting research, there is still need for continued analysis as some basic questions remain unresolved. Issues of highest priority include developing a better understanding of the microbial sources and sinks of these compounds and the organisms responsible for these processes. For example, a recent publication in Nature from Karl et al. (2008) demonstrates the aerobic production of methane at sea. There has long been an imbalance between the saturation state of the ocean with respect to methane and the atmosphere. The surface waters are supersaturated with respect to the atmosphere, which implies an unknown methane source and has consequently led to the perplexing ‘‘oceanic methane paradox.’’ Karl et al. (2008) provide evidence that this source is the aerobic production of biogenic methane. Traditionally, methane is often considered a product of anaerobic carbon remineralization, and while anaerobic degradation contributes the majority of biogenic methane to the Earth system, it is apparent that novel microbial processes and sources are still unknown. The ecological conditions and biogeochemistry of the organisms and environments of volatile hydrocarbon production are important areas of research. Research should build upon the foundation of previous discoveries in order to better understand the mechanisms and geochemical conditions responsible for the microbial production and consumption of other short-chain volatile hydrocarbons other than methane. In addition, fundamental questions regarding the evolution of these organisms and the Earth remain unresolved. Are there environmental and enzymatic relationships that preferentially lead to the production and consumption of these compounds? Are there specific evolutionary relationships between the environment and the enzymatic capability of these volatile hydrocarbon-forming organisms? Do these metal-based enzymes hint at or provide evidence of the conditions on early Earth, or throughout the evolution of the Earth? The biosynthesis of methane and non-CH4 hydrocarbons will continue to present an interesting avenue of research in the future, and there is much to be discovered. The microbial production and consumption of volatile hydrocarbons are important and unique processes that have far-reaching implications for all environments on Earth.

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Hattori S (2008) Syntrophic acetate-oxidizing microbes in methanogenic environments. Microbes Environ 23: 118–127. Hausrath EM, Liermann LJ, House CH, Ferry JG, Brantley SL (2007) The effect of methanogen growth on mineral substrates: will Ni markers of methanogen-based communities be detectable in the rock record? Geobiol 5: 49–61. Hinrichs KU, Hayes JM, Bach W, Spivack AJ, Hmelo LR, Holm NG, Johnson CG, Sylva SP (2006) Biological formation of ethane and propane in the deep marine subsurface. Proc Natl Acad Sci 103: 14684–14689. Hippe H, Caspari D, Fiebig K, Gottschalk G (1979) Utilization of trimethylamine and other n-methyl compounds for growth and methane formation by Methanosarcina barkeri. Proc Natl Acad Sci 76: 494–498. Horbach S, Salm H, Welle R (1993) Isoprenoid biosynthesis in bacteria: two different pathway. FEMS Microbiol Lett 111: 135–140. Hunt JM (1974) Hydrocarbon geochemistry of the Black Sea. In: Degens ET, Ross DA (eds) The Black Sea – Geology, Chemistry, and Biology, Am. Assoc. Pet. Geol. Tulsa, Oklahoma, pp. 499–504. Jenden PD, Kaplan IR (1986) Comparison of microbial gases from the Middle America Trench and Scripps Submarine Canyon: implications for the origin of natural gas. Appl. Geochem 1: 631–646. Jones WJ, Leigh JA, Mayer F, Woese CR, Wolfe RS (1983) Methanococcus jannaschii sp nov, and extremely thermophilic methanogens from a submarine hydrothermal vent. Arch Microbiol 136: 254–261. Karl DM, Beversdorf L, Bjo¨rkman KM, Church MJ, Martinez A, DeLong EF (2008) Aerobic production of methane in the sea. Nature 1: 473–478. Kida K, Shigematsu T, Kijima K, Numagchi M, Mochiniga Y, Abe N, Morimura S (2001) Influence of Ni2+ and Co2+ on methanogenic activity and amounts of coenzymes involved in methanogenesis. J Biosci Bioengin 91: 590–595. King GM, Klug MJ, Lovley DR (1983) Metabolism of acetate, methanol and methylated amines in intertidal sediments of Lowes Cove, Maine. Appl Environ Microbiol 45: 1848–1853. Knies J, Damm E, Gutt J, Mann U, Pinturier L (2004) Near-surface hydrocarbon anomalies in shelf sediments off Spitsbergen: Evidence for past seepage. Geochem Geophys Geosys 6: 1–14. Kurr M, Huber R, Konig H, Jannasch HW, Fricke H, Trincone A, Kristjansson JK, Stetter KO (1991) Methanopurus Kandlerim gen and sp represents a novel group of hyperthermophilic methanogens, growing at 110
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The Microbial Production of Methane and Other Volatile Hydrocarbons Kvenvolden KA, Redden GD (1980) Hydrocarbon gases in sediment of the shelf, slope, and basin of the Bering Sea. Geochim Cosmochim Acta 44: 1145–1150. Ladygina N, Dedyukhina EG, Vainshtein MB (2006) A review on microbial synthesis of hydrocarbons. Process Biochem 41: 1001–1014. Martens CS, Berner RA (1974) Methane production in the interstitial waters of sulfate-depleted marine sediments. Science 185: 1167–1169. Nakagawa F, Yoshida N, Nojiri Y, Makarov VN (2002) Production of methane from alasses in eastern Siberia: Implications from its 14C and stable isotopic compositions. Global Biogeochem Cycles 16: 14.1–14.15. Nottingham PM, Hungate RE (1969) Methanogenic fermentation of benzoate. J Bacteriol 98: 1170–1172. Oremland RS (1981) Microbial formation of ethane in anoxic estuarine sediments. Appl Environ Microbiol 42: 122–129. Oremland RS, Marsh LM, Polcin SP (1982a) Methane production and simultaneous sulfate reduction in anoxic salt marsh sediments. Nature 296: 143–145. Oremland RS, DesMarais DJ (1982b) Methanogenesis in Big Soda Lake, Nevada: an alkaline, moderately hypersaline desert lake. Appl Environ Microbiol 43: 462–468. Oremland RS, Polcin S (1982c) Methanogenesis and sulfate reduction – competitive and noncompetitive substrates in estuarine sediments. Appl Environ Microbiol 44: 1270–1276. Oremland RS (1988) Biogeochemistry of methanogenic bacteria. In: Zehnder AJB (ed.). Biology of Anaerobic Microorganisms. New York: Wiley, pp. 641–705. Oremland RS, Whiticar MJ, Strohmaier FE, Kiene RP (1988) Bacterial ethane formation from reduced, ethylated sulfur compounds in anoxic sediments. Geochim Cosmochim Acta 52: 1895–1904. Rashid MA, Vilks G (1977) Environmental controls of methane production in Holocene basins in eastern Canada. Org Geochem 1: 53–59. Roy R, Klu¨ber HD, Conrad R (1997) Early initiation of methane production in anoxic rice soil despite the presence of oxidants. FEMS Microbiol Ecol 24: 311–320. Rudd JWM, Hamilton RD, Campbell NER (1974) Measurements of the microbial oxidation of methane in lake water. Limnol Oceanog 19: 519–524. Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61: 262–280. Schoell M (1980) The hydrogen and carbon isotopic composition of methane from natural gases of various origins. Geochim Cosmochim Acta 44: 649–661. Schoell M (1983) Genetic characterization of natural gases. AAPG Bull 67: 2225–2238. Schoell M (1988) Origins of methane in the Earth. Chem Geol 71: 1–10.

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Scho¨nheit P, Brandis A, Thauer RK (1979) Ferredoxin degradation in growing Clostridium pasteurianum during periods of iron deprivation. Arch Microbiol 120: 73–76. Smith AM, Cook RJ (1974) Implications of ethylene production by bacteria for biological balance of soil. Nature 252: 703–705. Smith MR, Mah RA (1978) Grwoth and methanogenesis by Methanosarcina strain 227 on acetate and methanol. Appl Environ Microbiol 36: 870–879. Stadtman TC, Barker HA (1949) Studies on the methane fermentation. VII. Tracer experiments on the mechanism of methane formation. Arch Biochem 21: 256–264. Valentine DL, Boone DR (2000) Diversity of methanogens. In: Seckbach J (ed.). Journey to Diverse Microbial Worlds. New York: Kluwer, pp 289–302. Valentine DL, Chidthaisong A, Rice A, Reeburgh WS, Tyler SC (2004) Carbon and hydrogen isotope fractionation by moderately thermophilic methanogens. Geochim Cosmochim Acta 68: 1571–1590. Vogel TM, Oremland RS, Kvenvolden KA (1982) Lowtemperature formation of hydrocarbon gases in San Francisco Bay sediment. Chem Geolo 37:289–298. Wagner WP, Nemecek-Marshall M, Fall R (1999) Three distinct phases of isoprene formation during growth and sporulation of Bacillus subtilis. J Bacteriol 181: 4700–4703. Weimer PJ, Zeikus JG (1978) Acetate metabolism in Methanosarcina barkeri. J Bacteriol 137: 332–339. Whelhan JA, Craig H (1979) Methane and hydrogen in East Pacific Rise hydrothermal fluids. Geophys Res Lett 6: 829–831. Whelan JK, Hunt JM, Berman J (1980) Volatile C1-C7 organic compounds in surface sediments from Walvis Bay. Geochim Cosmochim Acta 44: 1767–1785. Whiticar MJ, Faber E, Schoell M (1986) Biogenic methane formation in marine and freshwater environments: CO2 reduction vs. acetate formation – Isotope evidence. Geochim Cosmochim Acta 50: 693–709. Whiticar MJ (1990) A geochemical perspective of natural gas and atmospheric methane. Org Geochem 16: 531–547. Whiticar MJ, Suess E (1990) Hydrothermal hydrocarbon gases in the sediments of King George Basin, Bransfield Strait, Antarctica. Appl Geochem 5: 135–147. Whiticar MJ (1999) Carbon and hydrogen isotope systematics of bacterial formation of oxidation of methane. Chem Geol 161: 291–314. Wilkins K (1996) Volatile metabolites from actinomycetes. Chemosphere 32: 1427–1434. Winfrey MR, Ward DM (1983) Substrates for sulphate reduction and methane production in intertidal sediments. Appl Environ Microbiol 45: 193–199. Winfrey MR, Zeikus JG (1977) Effect of sulfate on carbon and electron flow during microbial methanogenesis

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in fresh water sediments. Appl Environ Microbiol 33: 275–281. Zeikus JG, Winfrey MR (1976) Temperature limitation of methanogenesis in aquatic sediments. Appl Environ Microbiol 31: 99–107. Zeikus JG, Kerby R, Krycki JA (1985) Single carbon chemistry of acetogenic and methanogenic bacteria. Science 227: 1167–1173. Zinder SH, Brock TD (1978) Production of methane and carbon dioxide from methane thiol and dimethyl

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7 Isoprene, Isoprenoids and Sterols J. Harder Department of Microbiology, Max Planck Institute for Marine Microbiology, Bremen, Germany [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128 2 The Isoprenoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 128 3 Terpene Synthases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129 4 Isoprenoid Biomarker . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130

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Abstract: Over 30,000 isoprenoids are synthesized from central metabolites by enzymes of the mevalonate-pathway or the alternative pathway, isoprenyl transferases, terpene synthases and modifying enzymes. Little is known on the degradation pathways and their metabolites.

1

Introduction

Isoprenoids are natural compounds derived from isoprene (2-methyl-buta-1,3-diene). Over 30,000 different substances are known that derived from multimers of the five carbon compound (Breitmaier, 2006; Conolly and Hill, 1992), including essential oils of plants, quinones in Bacteria and animals, ether lipids of Archaea, hopanoids in Bacteria and sterols in animals.

2

The Isoprenoids

The key intermediate of isoprenoid biosynthesis, isopentenyl diphosphate, is formed in two alternative pathways, the 1-deoxy-D-xylulose 5-phosphate pathway in plant chloroplasts, algae and most Bacteria, and the mevalonate pathway in most eukaryotes, Archaea and some Bacteria (Boucher and Doolittle, 2000; Hunter, 2007). Over 500 million tons of isoprene are released per year into the atmosphere (Guenther et al., 1995), but the photochemical oxidation is fast, with a half-live of a few hours. Beside plants, some bacteria release isoprene (Shirk et al., 2002).

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Head-to-tail condensation of the five carbon-intermediates dimethylallyl diphosphate and isopentenyl diphosphate yields geranyl diphosphate, the precursor molecule of all monoterpenes (Dev, 1982). The wide diversity and chemistry of monoterpenes was summarized by Erman (1985a, b). Sequiterpenes originate from the addition of isopentenyl diphosphate to geranyl diphosphate yielding farnesyl diphosphate. An addition of isopentenyl diphosphate to farnesyl diphosphate provides geranylgeranyl diphosphate, the precursor of diterpenes. The diversity of diterpenes in the largest family of flowering plants, Asteraceae, has been summarized by Seaman (1989), large collections of structural data were assembled by Dev (1985) and Rahman (1990). The condensation of geranylgeranyl diphosphate with isopentenyl diphosphate provides geranylfarnesyl diphosphate, the precursor for sesterterpenes. A different condensation type yields triterpenes: the tail-to-tail condensation of two farnesyl diphosphates to squalene. From squalene a variety of a-, mono-, di-, tri, tetra- and pentatriterpenoids including steroids can be formed (Dev, 1989). Over 1,500 pentacyclic triterpenes were reviewed by Ahmad and Rahman (1994). Saponins are plant glycosides with a 27-carbon steroid or a 30-carbon triterpene, they are amphiphilic detergents (Hostettmann and Marston, 1995). The tail-to-tail condensation of two geranylgeranylpyrophosphates yields 16-trans-phytone, a carotenoid (Britton et al., 2004) and the precursor of tetraterpenes. Polyterpenes comprise all isoprenoids that consist of more then eight isoprene units and are synthesized via head-to-tail condensation (Swiezewska and Danikiewicz, 2005). Prenylation describes the modification of proteins with farnesyl or geranylgeranyl residues (McTaggart, 2006). A specialty of marine systems, especially algae, is the halogenation of isoprenoic compounds (Faulkner, 2001).

3

Terpene Synthases

Terpene synthases (cyclases) catalyze the transformation of the diphosphate precursor into individual mono-, sequi- and diterpenes (Bohlmann et al., 1998). Crystal structures of cyclases

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provided new insights into the catalytic pathways via carbocations (Lesburg et al., 1998). The first squalene cyclase crystal structure, a recombinant squalene-hopene cyclase from Alicyclobacillus acidocaldarius, supported a proton-induced carbocationic cyclization reaction (Wendt et al., 1997). Mechanistic aspects of terpene cyclases have been reviewed by Croteau (1987), Wendt et al. (2000) and Christianson (2006, 2008) indicating a tight control of the multistep cationic addition reaction to carbon-carbon double bonds yielding the diversity of isoprenoid ring systems.

4

Isoprenoid Biomarker

Isoprenoids have a wide range of physico-chemical properties, due to the low polarity of many compounds primarily proportional to the molecular weight. Isoprene is a liquid with a high gas pressure and a short half-life in the atmosphere. To the opposite, polyisoprene is relative resistant to degradation, including microbial transformation and mineralization (Warneke et al., 2007). The low solubility and degradability of mainly compounds with more than 20 carbon atoms contributes to an increase portion of these compounds in the organic matter that is buried. Together with the diversity of isoprenoid structures, it is the basis for the intensive utilization of isoprenoids as biomarker in organic geochemistry (Peters et al., 2007, 2008). The compounds in the sediment reflect the source, mainly the primary producers plants and algae, and the biotransformations and diagenetic transformations during sedimentation. The addition of hydrogen sulfide to desaturated systems yielding sulfur heterocycles and the formation of saturated and aromatic rings in terpenoid compounds are typical reactions of the diagenesis.

5

Research Needs

Isoprenoids are the structurally most diverse natural compounds. The presence in living organisms and in sediments is explored intensively. The pathways to the precursors are well studied and a number of crystal structures have provided insight into the intramolecular cyclization reactions leading to the diversity of isoprenoid ring structures. Biotransformations during the sedimentation process are of interest for microbiologists and organic geochemists.

References Ahmad VU, Rahman AU (1994) Pentacyclic triterpenoids. In Handbook of Natural Products Data, vol. 2. Amsterdam: Elsevier. Bohlmann J, Mayer-Gauen G, Croteau R (1998) Plant terpenoid synthases: molecular biology and phylogenetic analysis. Proc Natl Acad Sci USA 95: 4126–4133. Boucher Y, Doolittle WF (2000) The role of lateral gene transfer in the evolution of isoprenoid biosynthesis pathways. Mol Microbiol 37: 703–716. Breitmaier E (2006) Terpenes: Flavor, Fragrances, Pharmaca, Pheromones. Weinheim: Wiley.

Britton G, Liaaen-Jensen S, Pfander H (2004) Carotenoids Handbook. Basel: Birkha¨user. Christianson DW (2006) Structural biology and chemistry of the terpenoid cyclases. Chem Rev 106: 3412–3442. Christianson DW (2008) Unearthing the roots of the terpenome. Curr Opin Chem Biol 12: 141–150. Conolly JD, Hill RA (1992) Dictionary of Terpenoids. New York: Chapman and Hall. Croteau R (1987) Biosynthesis and catabolism of monoterpenoids. Chem Rev 87: 929–954. Dev S (1982) CRC Handbook of Terpenoids: Monoterpenoids, vols. 1–2. Boca Racon: CRC Press.

Isoprene, Isoprenoids and Sterols Dev S (1985) CRC Handbook of Terpenoids: Diterpenoids, vols. 1–4. Boca Racon: CRC Press. Dev S (1989) CRC Handbook of Terpenoids: Triterpenoids, vols. 1–2. Boca Raton: CRC Press. Erman WF (1985a) Chemistry of the Monoterpenes: An Encyclopedic Handbook. Part A. New York: Marcel Dekker. Erman WF (1985b) Chemistry of the Monoterpenes: An Encyclopedic Handbook. Part B. New York: Marcel Dekker. Faulkner DJ (2001) Marine natural products. Nat Prod Rep 18: 1–49. Guenther AC, Hewitt C, Erickson D, Fall R, Geron C, Graedel T, Harley P, Klinger L, Lerdau M, McKay W, Pierce T, Scholes B, Steinbrecher R, Tallamraju R, Taylor J, Zimmermann P (1995) A global model of natural volatile organic compound emissions. J Geophys Res 100: 8873–8892. Hostettmann K, Marston A (1995) Saponins. Cambridge, MA: Cambridge University Press. Hunter WN (2007) The non-mevalonate pathway of isoprenoid precursor biosynthesis. J Biol Chem 282: 21573–21577. Lesburg CA, Caruthers JM, Paschall CM, Christianson DW (1998) Managing and manipulating carbocations in biology: terpenoid cyclase structure and mechanism. Curr Opin Struct Biol 8: 695–703. McTaggart SJ (2006) Isoprenylated proteins. Cell Mol Life Sci 63: 255–267.

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Peters KE, Walters CC, Moldowan JM (2007) The Biomarker Guide, Vol. 1: Biomarkers and Isotopes in the Environment and Human History. Cambridge, MA: Cambridge University Press. Peters KE, Walters CC, Moldowan JM (2008) The Biomarker Guide, Vol. 2: Biomarkers and Isotopes in Petroleum Systems and Earth History. Cambridge, MA: Cambridge University Press. Rahman AU (1990) Diterpenoid and steroidal alkaloids. In Handbook of Natural Products Data, vol. 1. Amsterdam: Elsevier. Seaman F (1989) Diterpenes of Flowering Plants: Compositae (Asteracae). New York: Springer. Shirk MC, Wagner WP, Fall R (2002) Isoprene formation in Bacillus subtilis: a barometer of central carbon assimilation in a bioreactor? Biotechnol Prog 18: 1109–1115. Swiezewska E, Danikiewicz W (2005) Polyisoprenoids: structure, biosynthesis and function. Pro Lipid Res 44(4): 235–258. Warneke S, Arenskotter M, Tenberge KB, Steinbuchel A (2007) Bacterial degradation of poly(trans1,4-isoprene) (gutta percha). Microbiology 153: 347–356. Wendt KU, Poralla K, Schulz GE (1997) Structure and function of a squalene cyclase. Science 277: 1811–1815. Wendt KU, Schulz GE, Corey EJ, Liu DR (2000) Enzyme mechanisms for polycyclic triterpene formation. Angew Chem Int Ed 39: 2812–2833.

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8 Hopanoids M. Rohmer Institut Le Bel, Universite´ de Strasbourg, Strasbourg CNRS, France [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 134 2 Structures of Bacterial Hopanoids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 135 3 Hopanoid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136 4 Putative Role: Hopanoids as Membrane Stabilizers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137 5 Hopanoid Distribution in Eubacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 138 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139

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Abstract: Pentacyclic triterpenoids of the hopane series are widespread in the organic matter of nearly all sediments. They represent the molecular fossils of the C35 bacteriohopanepolyols, a long overlooked series of bacterial metabolites. Their structural diversity, biosynthesis, distribution among bacteria and ecological significance are discussed in this contribution.

1

Introduction

Pentacyclic triterpenes of the hopane series 1 (hopanoids, > Fig. 1) represent one of the most abundant natural product series (Ourisson et al., 1984). They are found in the organic matter of all sediments, whatever is their age, origin or nature (Ourisson and Albrecht, 1992). Such a widespread occurrence suggested that they represent the molecular fossils of ubiquitous living organisms. This hypothesis was supported by the fact that diploptene 2 (> Fig. 1), a C30 hopanoid, was detected in the early 1970’s in a few cyanobacteria (Gelpi et al., 1970), in the methanotrophic bacterium, Methylococcus capsulatus, (Bird et al., 1971) and the thermoacidophile Alicyclobacillus acidocaldarius (De Rosa et al., 1971) and proved later to be correct. With respect to the ubiquitous occurrence of the fossil hopanoids, bacteria were the right candidates. (> Chapter 11, Vol. 1, Part 2) The major compounds in the extant organisms are,

. Figure 1 Bacterial triterpenes of the hopane series.

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however, always the C35 hopanoids that proved to be widely distributed among eubacteria. Their discovery was fortuitous. The search for compounds responsible for the orientation of the cellulose microfibrils excreted by ‘‘Acetobacter xylinum’’ showed that the active fraction contained hopane derivatives with an additional C5 tetrol side-chain (Haigh et al., 1973; Fo¨rster et al., 1973).

2

Structures of Bacterial Hopanoids

Triterpenes can be divided into two groups, depending on the biosynthetic origin of their polycyclic skeleton. The most common pathway is the cyclization of oxidosqualene leading to triterpenes with an oxygenated function at C-3. The occurrence of C30 3b-hydroxy or 3oxohopanoids is restricted, however, to a few angiosperms. In contrast, the 3-deoxyhopanoids, which derive from the direct cyclization of squalene, are much more frequently encountered. Such C30 hopanoid hydrocarbons as well as their oxidation products are found in scattered taxa of lower eukaryotes, e.g., in a few fungi, some lichens, mosses and ferns. Most abundantly, the 3-deoxyhopanoids are found in bacteria. In all hopanoid producing bacteria, two C30 hopanoids, diploptene 2 and diplopterol 3 (> Fig. 1), are usually present. Tetrahymanol 4 derivatives with the quasi-hopanoid gammacerane skeleton are characteristic of Rhodopseudomonas palustris and the related Bradyrhizobium spp. (Bravo et al., 2001). The major compounds are always the C35 bacteriohopanepolyols, which are characterized by a polyfunctionalized C5 n-alkyl side chain linked by a carbon/carbon bond to the triterpene skeleton. Bacteriohopanetetrol 5, which was even reported from the 50 millions years old Messel Eocene oil shale (Mycke et al., 1987), and aminobacteriohopanetriol 6 are the most common representatives of the series (Rohmer, 1993). Structural variations found on the C35 skeleton include the presence of the two diastereomers at C-22, of additional hydroxy groups at C-31 and/or C-30, of a double bond at C-6, C-11 or in both positions or of an additional methyl group either at C-3b, C-2b, C-2a (Rohmer, 1993) or C-31 (Simonin et al., 1994). Additional variations are found on the moieties linked to the terminal functional group: aminohexoses (7, > Fig. 1), N-acylaminohexoses or hexosuronic acids (8, > Fig. 1) linked to the C-35 hydroxy group of tetrol 5 via a glycosidic bond (Langworthy et al., 1976; Llopiz et al., 1992), a carbapseudopentose (9, > Fig. 1) linked via an ether bond (Renoux and Rohmer, 1985), carbamoyl groups at C-35 and C-34 or amino-acids (tryptophane, as in hopanoid 10, or lysine) or fatty acids linked via an amide bond to the terminal amino group of aminotriol 6 (Rohmer, 1993; Seemann et al., 1999). The structures of adenosylhopane 11 (Neunlist and Rohmer, 1985) and of lactone 12 (Seemann et al., 1999) suggest that the side-chain may be derived from a D-ribose derivative. Nearly all biohopanoids are characterized by a 17b configuration. The 17a configuration was until recently only found in hopanoid molecular fossils. Their formation was interpreted in terms of isomerization of the least stable 17b-biohopanoids during maturation of the organic matter in sediments. 17a-Bacteriohopanetetrol 12 (> Fig. 1) was, however, unexpectedly detected in a few Gram-positive bacteria such as Frankia and related species (Rosa-Putra et al., 2001). In fact, 17a-hopanoids are probably more widely distributed among eubacteria, as suggested by their detection in modern sediments, e.g., 17a-bis-homohopanoic acid 14 with a degraded side chain in microbial mats from methane seeps in the Black Sea (Thiel et al., 2003) or 17a-bacteriohopanetetrol 13 in Holocene sediments from Ace Lake ‘‘Antarctica’’ (Talbot et al., 2000a).

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Because of this huge structural diversity resulting in compounds of quite different polarities, there is no general method for the screening of intact hopanoids. In addition, the native hopanoids are amphiphilic and insoluble in most usual solvents, two features making their handling uneasy. Isolation and characterization of all intact bacterial hopanoids was therefore performed on their peracetylated derivatives, which can be isolated by chromatographic methods and detected in the resulting fractions by the characteristic methyl singlet pattern observed on the 1H-NMR spectra. A general method for the detection of biohopanoids is the periodic acid oxidative cleavage of the polyhydroxylated side-chain, which affords aldehydes. A subsequent sodium borohydride reduction yields a mixture of primary alcohols that is easily purified and analyzed by GC-MS upon acetylation (Rohmer et al., 1984) or silylation. Although this reaction sequence results in a loss of information on the structural diversity of the biohopanoid side-chains, this approach is still the only general method, which allows the detection of most known bacterial hopanoids. Provided that reference material is available, the recently developed atmospheric pressure chemical ionization mass spectrometry coupled to liquid chromatography on the peracetylated derivatives represents an interesting alternative to be further investigated for hopanoid detection (Talbot et al., 2001, 2003a, b, 2008a).

3

Hopanoid Biosynthesis

First results on the biosynthesis of bacterial hopanoids were obtained on the formation of the pentacyclic ring system. Cell-free systems of the bacterium ‘‘Acetobacter pasteurianus’’ cyclized tritium-labeled squalene 15 into diploptene 2 and diplopterol 3 (> Fig. 2) (Anding et al., 1975). Lack of specificity of this system was pointed out. It cyclizes both enantiomers of 2,3-oxidosqualene, which is not the natural substrate of the enzyme: the (3S)-enantiomer as expected into 3b-hydroxyhopanoids, and the non-natural (3R)-enantiomer into 3ahydroxyhopanoids with different foldings of the two acyclic substrates (Bouvier et al., 1980). The squalene/hopene cyclase of Alicyclobacillus acidocaldarius has later been isolated, cloned, expressed and sequenced (Ochs et al., 1992). The corresponding gene was identified, and the availability of the recombinant protein led to the first tridimensional structure of a triterpene

. Figure 2 Cyclization of squalene 15 into diploptene 2 and diplopterol 3 by bacterial squalene/hopene cyclases.

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synthase (Wendt et al., 1997; Reinert et al., 2004), which permitted, by directed mutagenesis, the first extensive mechanistic studies on an enzyme catalyzing such a complex cyclization reaction (Feil et al., 1996; Hoshino and Sato, 2002). In order to find out the origin of the polyhydroxylated side-chain of the C35 bacteriohopane derivatives, labeling experiments, first with 13C labeled acetate and later with glucose isotopomers, were performed using selected hopanoid producing bacteria. The C5 side chain was thus shown to be derived from a D-ribose derivative linked by a carbon/carbon via its C-5 carbon atom to the isopropyl group of the hopane moiety (Flesch and Rohmer, 1988; Rohmer et al., 1989). This biogenetic pathway was already suggested by the structures of adenosylhopane 11 and lactone 12 (> Fig. 1). Genes and enzymes involved in this original alkylation process are yet unknown. More interesting, the labeling patterns found in the isoprene units of the triterpene moiety did not correspond to those expected from the mevalonate pathway, which was unanimously admitted as the sole biosynthetic pathway towards isopentenyl diphosphate and dimethylallyl diphosphate, the universal precursors of isoprenoids in living organisms (> Chapter 7, Vol. 1, Part 2). Extensive investigations using 13C and 2H labeled precursors led to the discovery of the mevalonate-independent methylerythritol phosphate pathway for the formation of isoprene units (Rohmer et al., 1993; Rohmer, 2007). This metabolic route is the major pathway for the biosynthesis of isoprenoids in bacteria, including hopanoids. It is also present in the plastids of all phototrophic organisms (plants, algae) and phylogenetically related taxa (e.g., the Plasmodium spp. responsible for malaria). The last biosynthetic step concerns the introduction of additional methyl groups, which are found on saturated carbon atoms of the hopane skeleton. Methionine is the initial methyl donor for the additional methyl groups of 3b-methylbacteriohopaneterols of ‘‘Acetobacter pasteurianus,’’ 2b-methyldiplopterol of Methylobacterium organophilum (Zundel and Rohmer, 1985) or 31-methylbacteriohopanetetrol of Acetobacter europaeus (Simonin et al., 1994). They were all labeled by an unknown methylation process upon feeding with [CD3] methionine with retention of all three deuterium atoms.

4

Putative Role: Hopanoids as Membrane Stabilizers

Apparently, hopanoids are essential metabolites for many hopanoid producers; inhibition of their biosynthesis by a squalene/hopene cyclase inhibitor, such as 2,3-dihydro-2-azasqualene, is lethal, whereas this inhibitor does not affect the growth of organisms not producing hopanoids (Flesch and Rohmer, 1987). Bacterial hopanoids, like cholesterol, are amphiphilic molecules with an apolar pentacyclic triterpene hydrocarbon skeleton and a hydrophilic polyhydroxylated side-chain. Like the sterols, the ring junctions are all trans, leading to a rigid and planar ring system optimized for interactions with the hydrocarbon chains of the phospholipid fatty acids. In addition, the length of the hydrophobic moiety roughly corresponds to half of the thickness of a membrane phospholipid bilayer. All these structural and biosynthetic features characterize possible membrane stabilizers and support the hypothesis that hopanoids in bacteria may represent the equivalents of the sterols found in the eukaryotic plasma membranes and were the phylogenetic precursors of sterols (Rohmer et al., 1979; Ourisson et al., 1987). The structural diversity of the side-chains as well as the modifications on the polycyclic ring system probably play a minor role as long as the structural characteristics required for a membrane stabilizers are fulfilled. This does not exclude other, yet unknown, roles for hopanoids in bacterial cells.

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In fact, in membrane models, hopanoids were shown to modulate the membrane stability and fluidity much like sterols do (Kannenberg and Poralla, 1980; Poralla et al., 1980; Benz et al., 1983; Bisseret et al., 1983; Chen et al., 1995a, b; Sato et al., 1995). In vivo, hopanoid concentration increased in Alicyclobacillus acidocaldarius with the temperature, counterbalancing the destabilizing effect of temperature on the membrane (Poralla et al., 1984). In Zymomonas mobilis, a bacterium tolerating high ethanol concentrations (up to 13%, wt/vol in batch cultures), the extremely high and constant hopanoid concentration (30 mg/g, dry weight) is believed to contribute to the ethanol resistance of the bacterium (Hermans et al., 1991). In a similar way, bacteriohopanetetrol derivatives are accumulated in large amounts in vesicles of the nitrogen fixing bacterium Frankia spp., constituting most probably a barrier to oxygen and participating in the protection of the oxygen sensitive nitrogenase (Berry et al., 1993). In Streptomyces coelicolor, like in all investigated Streptomyces spp., hopanoids are produced in minute amounts (ca. 2–4 mg/g, dry weight) in submersed cultures growing vegetatively. Hopanoid concentration is increased by a factor 1000 in emersed sporulating cultures. S. coelicolor mutants defective in the formation of aerial mycelium and spores do not produce hopanoids (Poralla et al., 2000). This observation suggests that hopanoids may fulfill other roles in bacterial cells, but very little information is available on this aspect. The few available observations suggest, however, that the production of hopanoids may confer some ecological advantages to the producers, especially in stress conditions (heat, pH, oxygen, solvents. . .).

5

Hopanoid Distribution in Eubacteria

First data concerning the distribution of hopanoids in prokaryotes were obtained by the H5IO6/NaBH4 side-chain derivatization method followed by GC-MS analysis of the resulting acetylated primary alcohol fractions (Rohmer et al., 1984). This first study pointed out to the broad distribution of the C35 bacteriohopane members of this triterpene series among eubacteria, whereas they are absent in eukaryotes and archaea. The choices of the first investigated strains were made at a time where structure elucidations represented a priority and led to the discovery of structural diversity of bacterial hopanoids. Although an as broad as possible coverage of the bacterial diversity was the goal of the studies, some genera (e.g., Acetobacter, Methylobacterium, methanotrophs) or groups (e.g., cyanobacteria), which were known as good hopanoid producers, were overrepresented in the screening lists, whereas strains presenting difficulties in their cultivation, for instance anaerobic bacteria, were underrepresented. This changed during the last years. Hopanoids were repeatedly reported from bacteria that are characteristic of anoxic environments (Thiel et al., 2003; Sinninghe Damste´ et al., 2004; Ha¨rtner et al., 2005; Fischer et al., 2005; Blumenberg et al., 2006; Rattray et al., 2008). Many publications later completed the first list, using mainly the same derivatization method, and more recently APCI-ion trap LCMS of the acetylated intact biohopanoids (Talbot et al., 2003a, b). Such chemical analyses are now efficiently completed by the search of the genes involved in the hopanoid biosynthesis. Screenings focuses always on the squalene/hopene cyclase (> Fig. 2) gene, which is the only known gene of this biosynthetic pathway. The results are most likely relevant for the presence of the whole biosynthetic pathway towards the C35 bacteriohopanepolyols. Indeed, no hopanoid producer is presently known, synthesizing only diploptene 2 and/or diplopterol 3 (> Fig. 2), the two reaction products of the squalene/hopene cyclase. Knowledge of the subsequent genes in the biosynthetic pathway leading to the major C35

Hopanoids

8

bacteriohopane derivatives would strengthen such investigations. Genome analysis reveals the capacity to produce or not hopanoids. Indeed, negative results obtained by the chemical analyses do not differentiate between the absence of the hopanoid biosynthetic pathway and the lack of its expression in an effective hopanoid producer (like Streptomyces coelicolor, which does not significantly synthesize hopanoid in vegetative growth conditions and highly expresses the pathway when sporulating). The presently known distribution of hopanoids in eubacteria shows that these metabolites are widely distributed in many groups. Although some groups were regularly found as hopanoid producers among Gram-Positive or Gram-negative (cyanobacteria, acetic acid bacteria, rhodospirillaceae, Rhodopseudomonas group, Streptomyces, Frankia. . .), no general rules can be drawn from the present knowledge. There is no dichotomy between aerobes and anaerobes, the hopanoid biosynthesis, in contrast to sterol biosynthesis, being completely oxygen independent. In addition, less than 1% of the bacteria species found in a natural environment can be detected by the usual methods of microbiology (Palleroni, 1997; Dykhuizen, 1998). Metagenomic analyses for the squalene/hopene cyclase gene has thus been performed on whole bacterial populations in order to circumvent this problem (Pearson et al., 2007). This method has for instance shown that hopanoid producing strains are apparently not numerous in lacustrine and marine environments.

6

Research Needs

Biomarkers of the hopane series are widely used in organic geochemistry. Many problems are, however, still linked to this natural product family. A large amount of knowledge has been gathered on the bacterial hopanoids since the first record of diploptene in a few prokaryotes in the early 1970s. It is probable that the structural diversity of this series has not been completely explored. Genome analyses (Pearson et al., 2007) as well as attempts to correlate in sediments preserved 16S rRNA genes with the likely source organisms for hopanoids (Coolen et al., 2008) will give a more accurate overview of their distribution among eubacteria in the future. In order to get the maximum of information, one should attempt to know the distribution of each individual structure of the intact hopanoids. This information might be helpful for deciphering the physiological role of these compounds and for the reconstitution of a palaeoenvironment from their molecular fossils record (Talbot and Farrimond, 2007). For instance, 2b-methylhopanoids are used as biomarkers for cyanobacteria, although their finding in a Rhodopseudomonas palustris strain challenges this possibly too narrow interpretation (Rashby et al., 2007), and 3b-methylhopanoids as well as the C35 aminopolyols represent the signature of obligate methanotrophs. Finally, very little is known about the reactions (biotic or abiotic) implied in the conversion of a bacterial biohopanoid into the geohopanoids found in sediment.

References Anding C, Rohmer M, Ourisson G (1975) Non-specific biosynthesis of hopane triterpenes in a cell-free system from Acetobacter rancens. J Am Chem Soc 98: 1274–1275.

Benz R, Hallmann, Poralla K, Eibl H (1983) Interaction of hopanoids with phosphatidylcholines containing oleic and o-cyclohexyldodecanoic acid in lipid bilayer membranes. Chem Phys Lipids 34: 7–24.

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Berry AM, Harriottt OT, Moreau RA, Osman SF, Benson DR, Jones AD (1993) Hopanoid lipids compose the Frankia vesicle envelope, presumptive barrier of oxygen diffusion to nitrogenase. Proc Natl Acad Sci USA 90: 6091–6094. Bird CW, Lynch JM, Pirt SJ, Reid WW (1971) The identification of hop-22(29)-ene in prokaryotic organisms. Tetrahedron Lett 34: 3189–3190. Bisseret P, Wolff G, Albrecht AM, Tanaka T, Nakatani Y, Ourisson G (1983) A direct study of the cohesion of lecithin bilayers: the effect of hopanoids and a,odihydroxycarotenoids. Biochem Biophys Res Commun 110: 320–324. Blumenberg M, Kru¨ger M, Neuhaus K, Talbot HM, Oppermann BI, Seifert R, Pape T, Michaelis W (2006) Biosynthesis of hopanoids by sulfatereducing bacteria (genus Desulfovibrio). Environm Microbiol 8: 1220–1227. Bouvier P, Berger Y, Rohmer M, Ourisson G (1980) Nonspecific gammacerane triterpene biosynthesis by cell-free system from the protozoon Tetrahymena pyriformis. Conformation of squalene, (3S)squalene epoxide and (3R)-squalene epoxide during the cyclization. Eur J Biochem 112: 557–560. Bravo JM, Perzl M, Ha¨rtner T, Kannenberg EL, Rohmer M (2001) Novel methylated triterpenoids of the gammacerane series from the nitrogen-fixing bacterium Bradyrhizobium japonicum USDA 110. Eur J Biochem. 268:1323–1331. Chen Z, Sato Y, Nakazawa I, Suzuki Y (1995a) Interactions between bacteriohopane-32,33,34,35-tetrol and liposomal membranes composed of dipalmitoylphosphatidylcholine. Biol Pharm Bull 18: 477–480. Chen Z, Tanno N, Takenaka S, Suzuki Y (1995b) Effects of bacteriohopane-32-ol on the stability of various kinds of liposomal membranes. Biol Pharm Bull 18: 600–604. Coolen MJL, Talbot HM, Abbas BA, Ward C, Schouten C, Volkman JK, Sinninghe Damste´ JS (2008) Sources of sedimentary bacteriohopanepolyols as revealed by 16S rDNA stratigraphy. Environm Microbiol 10: 1783–1803. De Rosa M, Gambacorta A, Minale L (1971) Bacterial triterpenes. Chem Commun 619–620. Dykhuizen DE (1998) Santa Rosalia revisited: why are there so many species of bacteria? Antonie van Leeuwenkoek 73: 25–33. Feil C, Su¨ssmuth R, Jung G, Poralla K (1996) Site-directed mutagenesis of putative active-site residues in squalene-hopene cyclase. Eur J Biochem 242: 51–55. Fischer WW, Summons RE, Pearson A (2005) Targeted genomic detection of biosynthetic pathways: anaerobic production of hopanoid biomarkers by a common sedimentary microbe. Geobiology 3: 33–40.

Flesch G, Rohmer M (1987) Growth inhibition of hopanoid synthesizing bacteria by squalene cyclase inhibitors. Arch Microbiol 147: 100–104. Flesch G, Rohmer M (1988) Prokaryotic triterpenoids. The biosynthesis of the bacteriohopane skeleton: formation of isoprenic units from two distinct acetate pools and a novel type of carbon/carbon linkage between a triterpene and D-ribose. Eur J Biochem 175: 405–411. Fo¨rster HJ, Biemann K, Haigh WG, Tattrie NH, Colvin JR (1973) The structure of novel C35 pentacyclic terpenes from Acetobacter xylinum. Biochem J 135: 133–143. Gelpi E, Schneider H, Mann J, Oro J (1970) Hydrocarbons of geochemical significance in microscopic algae. Phytochemistry 9: 603–612. Haigh WG, Fo¨rster HJ, Biemann K, Tattrie NH, Colvin JR (1973) Induction of orientation of bacterial cellulose microfibrils by a novel terpenoid from Acetobacter xylinum. Biochem J 135: 145–149. Ha¨rtner T, Straub KL, Kannenberg E (2005) Occurrence of hopanoid lipids in anaerobic Geobacter species. FEMS Microbiol Lett 243: 59–64. Hermans MAF, Neuss B, Sahm H (1991) Content and composition of hopanoids in Zymomonas mobilis under various growth conditions. J Bacteriol 173: 5592–5595. Hoshino T, Sato T (2002) Squalene-hopene cyclase: catalytic mechanisms and substrate recognition. J Chem Soc Chem Commun 291–301. Kannenberg E, Poralla K (1980) A hopanoid from the thermo-acidophilic Bacillus acidocaldarius condenses membranes. Naturwissenschaften 67: 458–459. Langworthy TA, Mayberry WR, Smith PF (1976) A sulfonolipid and novel glucoamidyl glycolipid from the extreme thermoacidophile Bacillus acidocaldarius. Biochim Biophys Acta 431: 550–569. Llopiz P, Neunlist S, Rohmer M (1992) Prokaryotic triterpenoids. O-a-D-glucuronopyranosylbacteriohopanetetrol, a novel hopanoid from the bacterium Rhodospirillum rubrum. Biochem J 287: 159–162. Mycke B, Narjes F, Michaelis W (1987) Bacteriohopanetetrol from chemical degradation of an oil shale kerogen. Nature 326: 179–181. Neunlist S, Rohmer M (1985) A novel hopanoid, 30-(50 adenosyl)hopane from the purple non sulfur bacterium Rhodopseudomonas acidophila with possible DNA interactions. Biochem J 228: 769–771. Ochs D, Kaletta C, Entian KD, Beck-Sickinger A, Poralla K (1992) Cloning, expression and sequencing of squalene-hopene cyclase, a key-enzyme in triterpenoid metabolism. J Bacteriol 174: 298–302. Ourisson G, Albrecht P (1992) Hopanoids. 1. Geohopanoids: the most abundant natural products on earth? Acc Chem Res 25: 398–402.

Hopanoids Ourisson G, Albrecht P, Rohmer M (1984) The microbial origin of fossil fuels. Scientific American 251: 44–51. Ourisson G, Rohmer M, Poralla K (1987) Prokaryotic hopanoids and other polyterpenoid sterol surrogates. Annu Rev Microbiol 41: 301–333. Palleroni NJ (1997) Prokaryotic diversity and the importance of culturing. Antonie van Leeuwenhoek 72: 3–19. Pearson A, Flood Page SR, Jorgenson TL, Fischer WW, Higgins MB (2007) Novel hopanoid cyclases in the environment. Environm Microbiol 9: 2175–2188. Poralla K, Ha¨rtner T, Kannenberg E (1984) Effect of temperature and pH on the hopanoid content of Bacillus acidocaldarius. FEMS Microbiol Lett 23: 253–256. Poralla K, Kannenberg E, Blume A (1980) A glycolipid containing hopane isolated from the acidophilic thermophilic Bacillus acidocaldarius has a cholesterol-like function in membranes. FEBS Lett 113: 107–110. Poralla K, Muth G, Ha¨rtner T (2000) Hopanoids are formed during transition from substrate to aerial hyphae in Streptomyces coelicolor A3(2). FEMS Microbiol Lett 189: 93–95. Rashby SE, Sessions AL, Summons RE, Newman DK (2007) Biosynthesis of 2-methylhopanoids by an anoxygenic phototroph. Proc Natl Acad Sci USA 104: 15099–15104. Rattray JE, van de Vossenberg J, Hopmans EC, Kartal B, van Niftrik L, Rijpstra WIC, Strous M, Jetten MSM, Schouten S, Sinninghe Damste´ J (2008) Ladderane lipid distribution in four genera of anammox bacteria. Arch Microbiol 190: 51–66. Reinert DJ, Balliano G, Schulz GE (2004) Conversion of squalene to the pentacyclic hopene. Chem Biol 11: 121–126. Renoux JM, Rohmer M (1985) Prokaryotic triterpenoids. New bacteriohopanetetrol cyclitol ethers from the methylotrophic bacterium Methylobacterium organophilum. Eur J Biochem 151: 405–410. Rohmer M, Bouvier P, Ourisson G (1979) Molecular evolution of biomembranes: structural equivalents and phylogenetic precursors of sterols. Proc Natl Acad Sci USA 76: 847–851. Rohmer M (1993) The biosynthesis of triterpenoids of the hopane series in Eubacteria: a mine of new enzymes reactions. Pure Appl Chem 65: 5295–5298. Rohmer M (2007) Diversity in isoprene unit biosynthesis: the methylerythritol phosphate pathway in bacteria and plastids. Pure Appl Chem 79: 739–751. Rohmer M, Bouvier-Nave´ P, Ourisson G (1984) Distribution of hopanoid triterpenes in prokaryotes. J Gen Microbiol 130: 1137–1150.

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Rohmer M, Knani M, Simonin P, Sutter B, Sahm H (1993) Isoprenoid biosynthesis in bacteria: a novel pathway for the early steps leading to isopentenyl diphosphate. Eur J Biochem 295: 517–524. Rohmer M, Sutter B, Sahm H (1989) Bacterial sterol surrogates. Biosynthesis of the side-chain of bacteriohopanetetrol and of a carbocyclic pseudopentose from 13C-labelled glucose in Zymomonas mobilis. J Chem Soc Chem Commun 19: 1471–1472. Rosa-Putra S, Nalin R, Domenach AM, Rohmer M (2001) Novel hopanoids from Frankia spp. and related soil bacteria. Squalene cyclization and significance of geological biomarkers revisited. Eur J Biochem 268: 4300–4306. Sato Y, Chen Z, Suzuki Y (1995) Thermodynamic effects of hopanoids on synthetic and bacterial phospholipid membranes. Chem Pharm Bull 43: 1241–1244. Seemann M, Bisseret P, Tritz JP, Hooper AB, Rohmer M (1999) Novel bacterial triterpenoids of the hopane series from Nitrosomonas europaea and their significance for the formation of the C35 bacteriohopane skeleton. Tetrahedron Lett 40: 1681–1684. Simonin P, Tindall B, Rohmer M (1994) Structure elucidation and biosynthesis of 31-methylhopanoids from Acetobacter europaeus: a new series of bacterial triterpenoids. Eur J Biochem: 22: 765–771. Sinninghe Damste´ J, Rijpstra WIC, Schouten S, Fuerst JA, Jetten MSM, Strous M (2004) The occurrence of hopanoids in planctomycetes: implications for the sedimentary biomarker record. Org Geochem 35: 561–566. Talbot HM, Farrimond P (2007) Bacterial populations recorded in diverse sedimentary biohopanoid distributions. Org Geochem 38: 1212–1225. Talbot HM, Marco JL, Sinninghe Damste´ J (2008a) An unusual 17a,21b(H)-bacteriohopanetetrol in Holocene sediments from Ace Lake (Antarctica). Org Geochem 39: 1029–1032. Talbot HM, Squier AH, Keely BJ, Farrimond P (2003b) Atmospheric pressure chemical ionization reversedphase liquid chromatography/ion trap mass spectrometry of intact bacteriohopanepolyols. Rapid Commun Mass Spectrom 17: 728–737. Talbot HM, Summons RE, Jahnke LL, Cockell CS, Rohmer M, Farrimond P (2008b) Cyanobacterial bacteriohopanepolyol signatures from cultures and natural environmental settings. Org Geochem 39: 232–263. Talbot HM, Summons R, Jahnke L, Farrimond P (2003a) Characteristic fragmentation of bacteriohopanepolyols during atmospheric pressure chemical ionization liquid-chromatography/ion trap mass spectrometry of intact bacteriohopanepolyols. Rapid Commun Mass Spectrom 17: 2788–2796.

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Talbot HM, Watson DF, Murrell JC, Carter JF, Farrimond P (2001) Analysis of intact bacteriohopanepolyols by reversed phase high-performance liquid chromatography-atmospheric pressure chemical ionization mass spectrometry. J Chromatogr A 921: 175–185. Thiel V, Blumenberg M, Pape T, Seifert R, Michaelis W (2003) Unexpected occurrence of hopanoids at gas seeps in the Black Sea. Org Geochem 34: 81–87.

Wendt KU, Poralla K, Schulz GE (1997) Structure and function of a squalene cyclase. Science 277: 1811–1815. Zundel M, Rohmer M (1985) Prokaryotic triterpenoids. 3. The biosynthesis of 2b-methylhopanoids and 3b-methylhopanoids of Methylobacterium organophilum and Acetobacter pasteurianus. Eur J Biochem 150: 35–39.

9 Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C A. Pearson Harvard University, Department of Earth and Planetary Sciences, Cambridge, MA, USA [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 144

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3 3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.1.5 3.2 3.2.1 3.2.2 3.2.3

Metabolic Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146 Life on C1 Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146 Calvin Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 146 rTCA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 148 Homoacetogenesis and CO2-Reducing Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . 150 Aerobic Methanotrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 Anaerobic Methanotrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 151 Heterotrophy and Fermentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Glycolysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Oxidation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 152 Aceticlastic Methanogenesis and Acetotrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153

4

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153

C Systematics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 145

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_9, # Springer-Verlag Berlin Heidelberg, 2010

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Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C

Abstract: Hydrocarbons utilized as metabolic substrates by microorganisms derive from a variety of sources. These many sources are imprinted with a range of initial values of d13C, reflecting the diverse biochemical pathways of inorganic carbon fixation to biomass. Subsequent recycling of organic matter, and especially processes involving C1 and C2 compounds such as methane and acetate, impart additional fractionation to carbon-containing substrates and products. The range of observed values of d13C imposed by these transformations is reviewed in this chapter, with emphasis on processes involving microorganisms.

1

Introduction

Total organic matter (TOM) is a heterogeneous mixture of material both produced and recycled by biological processes. The distribution of stable isotopes of carbon (12C and 13C) throughout global organic carbon reservoirs reflects the multiple pathways of carbon assimilation used by organisms living in each of these environments, as well as mass transfer among the reservoirs. For a classic description of the global carbon cycle, see Hedges (1992). The majority of primary production on land and in the oceans is mediated by organisms that employ oxygenic photosynthesis. In oceanic environments, most organic matter is believed to originate from phytoplanktonic biomass and subsequent heterotrophic degradation of this material, including significant bacterial production. Both open-ocean and continental margin sediments, however, also contain organic matter from other sources. Some fraction of TOM may be derived from land plant debris and terrigenous soils delivered by erosion. Other carbon may come from weathered continental shales or ancient kerogen. On land, organic carbon reservoirs include living plant biomass, soils, and water-saturated (often anaerobic) wetlands, peat bogs, and (frozen) arctic tundra. In the ocean, active hydrothermal systems, as well as cold seeps and methane hydrates, also are significant sources of organic matter. Although carbon fixed by primary producers is recycled almost quantitatively (> 99.9%) back into carbon dioxide and water (Hedges and Keil, 1995), a small fraction of organic matter escapes aerobic respiration and accumulates. Molecules that are especially recalcitrant, such as pigments, lipids and many structural macro-molecules, become concentrated (Tegelaar et al., 1989). Under reducing conditions, the remaining organic matter is degraded further by anaerobic heterotrophic organisms such as sulfate reducers, fermenters and methanogens. These alterations are referred to collectively as diagenesis. Kerogen, an amorphous structural network of biochemical subunits, forms from polymerization and crosslinking of residual biomolecules during diagenesis (e.g., Derenne et al., 1991; de Leeuw & Largeau, 1993). During these processes, lipids deposited with the original organic matter undergo cracking, isomerization, reduction, and aromatization, eventually losing all of their functional groups. Subsequently, with increasing burial temperature and pressure, thermal degradation generates bitumen and natural gas. Bitumen is defined as the hydrocarbon-rich fraction of organic matter that can be extracted from sediments and sedimentary rocks using organic solvents. Petroleum reservoirs are merely concentrated deposits of bitumenous cracking products of biogenic organic matter. By the time organic deposits reach burial temperatures exceeding 150–250
C, most residual gas and liquid hydrocarbons have been expelled and the kerogen is largely dehydrogenated to aromatic or graphitic carbon. At all stages of diagenesis and catagenesis, the produced hydrocarbons and methane can serve as substrates for microbes.

Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C

9

Diverse and abundant microbes have been found to inhabit deep sediments (Parkes et al., 1994), petroleum reservoirs (Head et al., 2003; Voordouw et al., 1996), and hydrothermal and geothermal systems (Chapelle et al., 2002; Copeland, 1936; Jannasch and Wirsen, 1979; Kelley et al., 2005). Because they can obtain both carbon and energy from the liquid and gaseous hydrocarbons found in these environments, carbon isotopic analysis of sedimentary microbes and their organic substrates can be used as a tool to reveal likely metabolic pathways of carbon assimilation and new processes (e.g., Hinrichs et al., 1999), and to help infer ancient microbial ecosystems and their sources of carbon (e.g., c.f., Hayes, 1993; Hinrichs, 2002).

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13

C Systematics

Carbon in the geosphere is composed of three naturally occurring isotopes, the stable isotopes 12 C (0.989) and 13C (0.011), and the cosmogenic radionuclide, 14C (1012). The distribution of the minor isotopes of carbon is affected by thermodynamic and kinetic fractionation processes (13C and 14C), and by radioactive decay (14C). The half-life of 14C is 5730 years, thus it can yield useful radiocarbon chronologies only over the most recent few tens of thousands of years. 14C is detectable in petroleum hydrocarbons and the microbes that consume them only if the petroleum is actively forming in modern systems (e.g., Guaymas Basin sediments; Pearson et al., 2005; Peter et al., 1991). Most isotopic analyses of organic matter focus primarily on the two stable isotopes of carbon. Thermodynamic fractionation governs the distribution of 13C and 12C between reservoirs whose carbon exchange is slow enough to approach isotopic equilibrium, and typically the associated isotope effects are small. An example is the speciation of dissolved inorganic carbon (DIC) in seawater: CO2(aq) , HCO3 , CO32. These equilibrium exchange reactions result in unequal isotope distribution between the dissolved species. Kinetic fractionation is the more important control on the 13C distribution during biologicallymediated carbon transfers. Most biochemical reactions that interconvert C1 compounds (CO2 or CH4) and biomass have large fractionation effects associated specifically with the binding between the substrate and its enzymatic catalyst. In contrast, most heterotrophic reactions involving complex organic substrates have small fractionations. Isotopic composition is expressed as the ratio of 13C to 12C in the substance, relative to the ratio in a standard material. The units of fractionation are parts per thousand, or ‘‘permil’’ (‰) according to formula (1) and the standard reference material is a carbonate rock (VPDB; Vienna Pee Dee Belemnite), which by definition has a d13CVPDB value of 0:   RA d13 C ¼  1  1000 ð1Þ RVPDB where R  13C/12C. The isotopic ratios of two chemical species are used to calculate the fractionation factor (2) between substrate and product in a biological reaction A!B, and this commonly is expressed as the fractionation effect, epsilon (3): RA ð1000 þ dA Þ  aA=B ¼ RB ð1000 þ dB Þ

ð2Þ


eA=B  aA=B  1  1000

ð3Þ

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Because 13C preferentially remains in A, in unidirectional reactions (such as biological cabon fixation) eA-B almost always is a positive number and products are ‘‘lighter’’ than reactants. Often the comparison of interest is simply the difference between two measured d13C values (4). In general, when the isotopic difference between chemical species A and B is small (< 30‰), the difference between D and e is negligible (< 1‰) and can be ignored. A more complete discussion of isotope ratio calculations can be found in Hayes (1993, 2001). DA=B  dA  dB  eA=B

ð4Þ

The distribution of 13C throughout the global inorganic and organic carbon reservoirs is governed by a combination of the fractionation processes mentioned above. Typical thermodynamic fractionation effects are in the range of 1–10‰ for inorganic processes (> Table 1). Typical kinetic isotope effects for biochemical processes are larger, usually tens of permil units. The enzymes utilized by organisms to fix carbon are categorized most easily by the metabolic pathways in which they are used and by the species of inorganic carbon substrate for which they are specific (> Table 1; > Fig. 1). Data in this table are presented for reactions at 25
C, except in the case of data from cultured species of thermophiles. Indeed, in geothermal reservoirs temperatures may exceed the conditions reported in > Table 1. Values of aA-B generally decrease at higher temperatures, resulting in smaller isotopic differences between reactants and products (Whiticar, 1999). Either inorganic or organic carbon molecules can serve as substrates for microorganisms. During inorganic carbon fixation, organisms utilizing pathways specific for HCO3 necessarily begin with a substrate that is heavier isotopically (more positive value of d13C) than organisms that fix CO2 directly (> Table 1; > Fig. 1a), while organisms that produce biomass from assimilated CH4 have more negative values of d13C (> Table 1; > Fig. 1b). The currently available information therefore can be divided into pathways that typically yield isotopically ‘‘heavy’’ biomass (rTCA, 3-HP; 3-HP/4-HB; Di/4-HB); pathways yielding isotopically intermediate biomass (Calvin-Benson cycle autotrophs, some methanogens, most heterotrophs); and pathways yielding isotopically light biomass (homoacetogens, some CO2-reducing methanogens, methylotrophic methanogens, all methanotrophs). Biomass from organisms expressing these varied pathways has the potential to accumulate in the long-term sedimentary record. This necessitates an understanding of the magnitude and range of values of d13C that could be observed in a variety of geologic - including sedimentary and geothermal - settings. The fractionations associated with these pathways are summarized below and in > Table 1.

3

Metabolic Pathways

3.1

Life on C1 Compounds

3.1.1

Calvin Cycle

In the Calvin-Benson-Bassham (CBB) cycle (Bassham et al., 1950), one molecule of CO2(g) is attached to the carrier molecule ribulose-1,5-bisphosphate (RuBP) to form a C6 sugar that quickly is processed to other intracellular intermediates. Fractionation of isotopes occurs during the initial carboxylation step, and the fractionation effect (e) observed between CO2(g) and the attached carboxyl (RuBP–*CO2) is 29‰ for Type Ib RuBisCO (Ribulose-1,5bisphosphate carboxylase/oxygenase; Roeske & O’Leary, 1984). All eukaryotic primary

Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C

9

. Table 1 Stable carbon isotope fractionation at 25
C, shown as equilibrium fractionation factors (aA/B) for inorganic processes and kinetic isotope effects («A/B) for unidirectional processes Equilibrium reactions

a (‰)

Reference

CO2(aq) , CO2(g)

0.0011

Raven, 1997

CO2(aq) + H2O , HCO3 + H+

0.0090

Raven, 1997

CO2(aq) , CaCO3(s, calcite)

0.0096

Raven, 1997

Fractionation by Enzymes

Substrate

RuBisCO (Type IA, prochlorophyte)

CO2

24

Scott et al., 2007

RuBisCO (Type IB, eukaryotic)

CO2

29

Roeske & O’Leary, 1984

RuBisCO (Type IB, cyanobacterial)

CO2

22

Guy et al., 1993

RuBisCO (Type II, proteobacterial)

CO2

18–23

Roeske & O’Leary 1985; Robinson et al., 2003

PEPC (relative to CO2(aq))

HCO3

Reference

«CO2-product (‰)

2–4

O’Leary et al., 1981

Pyruvate synthase

CO2

n/a

a-Ketoglutarate synthase

CO2

n/a

Isocitrate dehydrogenase

CO2

n/a

Acetyl-CoA carboxylase

HCO3 HCO3 HCO3

n/a

CODH, acetogenesis

CO2

59

Gelwicks et al., 1989

CODH, CO2-reducing methanogenesis

CO2

30–80

Gelwicks et al., 1994; Botz et al., 1996

CODH, aceticlastic methanogenesis

CH3COOH 21

Methane monooxygenase

CH4

Propionyl-CoA carboxylase Malonyl-CoA carboxylase

Metabolic Pathway Fractionations

n/a n/a

35

Gelwicks et al., 1994 Templeton et al., 2006 Reference

«substrate-biomass (‰)

Calvin-Benson-Bassham, Type I RuBisCO

12–26

a

Calvin-Benson-Bassham, Type II RuBisCO

5–25

a

rTCAb

2–13

a

3-HP

7

a

; Van der Meer et al., 2001

3-HP/4-HBb

0–8

a

Di/4-HBb

2–4

a

15–29

a

7–36

a

b,c

Acetogens CO2-reducing methanogens Aceticlastic methanogens Aerobic methanotrophs

0–5 16–31

; Takigiku, 1987

Londry et al., 2008 Summons et al., 1994; Templeton et al., 2006

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. Table 1 (Continued) Metabolic Pathway Fractionations

«substrate-biomass (‰)

Methylotrophic methanogens

20

Anaerobic methanotrophsd Heterotrophy (> C2)

0–40 1 to 2

Reference Summons et al., 1998 Orphan et al., 2002 DeNiro & Epstein, 1978; Blair et al., 1985

a Many original citations for the fractionation associated with metabolic pathways appear in House et al. (2003); those that do not, or that post-date 2003, are listed individually b Early reports of e values for Archaea (e.g., Holo & Sireva˚g, 1986; House et al., 2003) were based on the assumption that these taxa used rTCA or 3-HP, as 3-HP/4-HB and Di/4-HB had not yet been discovered. Data from House et al. have been subdivided here based on probable metabolic and taxonomic correlations discussed in Berg et al. (2007) and Huber et al. (2008) c With the discovery of 3-HP/4-HB in Archaea, it is now probable that the only known value of e for a true 3-HP species is that for Chloroflexus aurantiacus d None of the Archaea that are putative anaerobic oxidizes of methane (ANMEs) have been grown in pure culture; values of e are probable based on cells isolated from sediments

producers (algae and plants) fix carbon using type I enzymes, as do cyanobacteria. A second form of RuBisCO, known as Type II, is used by some Proteobacteria to fix CO2, although many Proteobacteria simultaneously carry the Type I form. Other RuBisCOs, including Type III of methanogens, and Type IV RuBisCO-like proteins, are not involved in the primary pathways of carbon fixation for their respective organisms (for recent reviews, see Badger and Bek, 2008, and Tabita, 2008). Carbon isotopes generally are fractionated less by Type II than by Type I enzymes (> Table 1). Intracellular processes, anaplerotic reactions, diffusive limitation, and the potential for carbon concentrating mechanisms (CCMs) and/or active import of HCO3 all can complicate the interpretation of d13C values of biomass produced by Calvin cycle autotrophs. These processes lead to expressed fractionations for overall metabolic pathways that are not equivalent to the fractionation factors measured for pure enzymes (> Table 1). Nevertheless, the vast majority of total global primary production is mediated by the CBB cycle. Carbon fixed by organisms employing the Type I version accounts for most of this organic matter and has an average value of d13C of 25‰. This carbon in turn becomes the substrate for subsequent diagenetic reactions, and ultimately for petroleum and gas generation. Accordingly, most petroleum has a value of d13C around 27‰.

3.1.2

rTCA

Some microbes living at hydrothermal vents and/or under anaerobic conditions employ an alternative pathway for carbon fixation known as the reverse tricarboxylic acid cycle (rTCA; Evans et al., 1966). In rTCA, all carbon transformations of the Krebs respiratory cycle are operated in reverse to fix CO2, with the important addition of an enzyme specific to rTCA, ATP-citrate lyase (> Fig. 1a). The resulting citrate is cleaved to release a C2 compound, acetylCoA, which is then carboxylated to yield the metabolic intermediate, pyruvate. Organisms using this pathway of carbon fixation include green sulfur bacteria (Chlorobiaceae), some

. Figure 1 Carbon assimilation pathways used by microorganisms, showing enzymes involved in carbon incorporation and/or steps imparting isotopic fractionation (blue), other key enzymes (green), and substrates (red). In all cases, the final product must be a C3 intermediate (e.g., pyruvate) that can be used for intracellular biosynthetic reactions. For more detailed descriptions, see White (2000), Strauss & Fuchs (1993), Berg et al. (2007), Kniemeyer et al. (2007), Moran et al. (2008), Huber et al. (2008) Bott and Thauer (1989). Abbreviations: PEP, phosphoenolpyruvate; OAA, oxaloacetate; PGA, phosphoglycerate; aKG, a-ketoglutarate. (a) Pathways using inorganic carbon substrates; (b) pathways using organic carbon substrates and CH4.

Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C

9 149

150

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Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C

hydrogen-oxidizing bacteria, and numerous Archaea (for citations, see House et al., 2003). In particular, organic matter generated chemosynthetically at marine hydrothermal vents may derive dominantly from rTCA and thus may constitute one of the few present-day ecosystems in which significant carbon fixation derives from a pathway other than the CBB pathway (Campbell et al., 2003). Fractionation of carbon isotopes generally is smaller in the rTCA pathway than in CBB. This would appear to be due to a more prominent role for enzymes such as phosphoenolpyruvate carboxylase (PEPC), which is specific for HCO3, rather than CO2, as a substrate (> Table 1, > Fig. 1a). However, in some respects the small fractionations exhibited by rTCA species remain a mystery. Organisms using this pathway have an overall isotope discrimination of 2–13‰ relative to their source CO2 (> Table 1), similar to organisms using 3-HP and the other hydroxyacid pathways (below). The small fractionation associated with rTCA arises despite the fact that every C3 unit apparently is synthesized from three enzymes specific for CO2, each of which could have a probable maximum value of e not dissimilar from that observed for RuBisCOs. Small expressed fractionations reflect the fact that the kinetic isotope effects for respiratory carboxylases – as for anaplerotic reactions – are much smaller than for RuBisCOs. Additional metabolic flux investigations of rTCA organisms are necessary to examine these processes. 3.1.2.1

3-HP and Variants

Recent discoveries have revealed several pathways of carbon fixation in which a 3- and/or 4-carbon hydroxyacid serve as central intermediates (> Fig. 1a). These pathways are known as 3-hydroxypropionate (3-HP; Strauss and Fuchs, 1993), 3-HP/4-hydroxybutyrate (3-HP/ 4-HB; Berg et al., 2007), and dicarboxylate/4-HB (Di/4-HB; Huber et al., 2008). Few carbon isotope measurements have been reported, but the available data suggest that fractionation between substrate and biomass is smaller even than that observed for rTCA (e 0–7‰; > Table 1). The magnitude of fractionation is consistent with the prevalence of enzymes that require HCO3 in the Bacteria and Archaea that employ the hydroxyacid pathways (Holo and Sireva˚g, 1986; House et al., 2003). In most cases, the reporting of isotopic data for organisms using these pathways pre-dates the discovery of the pathways themselves; often the organisms were proposed initially to be rTCA species. The data from early references have therefore been reassigned according to modern phylogenetic understanding of the distribution (Berg et al., 2007) of the hydroxyacid pathways (> Table 1). These pathways appear to be particularly important in non-methanogenic Archaea, including aerobic Crenarchaeota. Biomass of globally significant archaeal populations such as autotrophic members of the marine group I Crenarchaeota (Karner et al., 2001) very likely are more enriched in 13C than average primary production. This is an important consideration for interpreting past marine depositional conditions in times when Archaea may have been more abundant (Kupyers et al., 2001).

3.1.3

Homoacetogenesis and CO2-Reducing Methanogenesis

The biomass of homoacetogens and CO2-reducing methanogens has a 13C content that results from production of cellular material using the Acetyl-CoA pathway (> Fig. 1a). Values of e ranging from 7–36‰ relative to source CO2 have been measured for the homoacetogen Acetobacterium woodii (Preub et al., 1989), and from a variety of methanogenic Euryarchaeota (c.f., House et al., 2003; > Table 1). Values of e in this range yield biomass with values of d13C

Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C

9

similar to CBB organisms, although the range is larger for the Acetyl-CoA species. Notably, these fractionations apply to the biomass produced, not to the resulting catabolic products. In the case of homoacetogenesis, the product acetate is 50–60‰ depleted in 13C relative to the starting CO2, while in methanogenesis, the CH4 produced is 30–80‰ depleted in 13C; i.e., the fractionation expressed between CO2-product is greater than between CO2-biomass. The byproducts CH3COOH and CH4 are important substrates for other sedimentary microbes, thus ultimately influencing the values of d13C at the next trophic level, i.e., acetate oxidizers and methanotrophs, respectively.

3.1.4

Aerobic Methanotrophy

Aerobic methanotrophs (> Fig. 1b) oxidize CH4 to the initial intermediate, CH3OH, using either a particulate or a soluble form of methane monooxygenase (pMMO, sMMO). The biomass that forms when CH4 is the primary substrate shows considerable range of variability in its value of d13C, with final fractionation relative to the source methane generally 16–31‰, although in conditions of CH4 limitation, the fractionation can be considerably less (Summons et al., 1994). Recent work suggests these effects – as well as differences that have been noted between sub-sets of aerobic methanotrophs known as Type I, Type II, and Type X methanotrophs – result from changes in the intracellular partitioning of carbon and are not the result of different kinetic isotope effects for pMMO and sMMO (Templeton et al., 2006). Because values of d13C of thermogenic and biogenic methane usually are < 40‰ for the former, and < 60‰ for the latter (Whiticar, 1999), organic matter formed by aerobic methanotrophs should have final values of d13C < 60 to 80‰.

3.1.5

Anaerobic Methanotrophy

A similar spectrum of very negative values of d13C is observed in the biomass of Archaea that are putative anaerobic methanotrophs (ANME species; Orphan et al., 2001, Orphan et al., 2002). It was proposed recently that the mechanism for the anaerobic oxidation of methane (AOM) proceeds via production of intermediate methyl sulfides (Moran et al., 2007; > Fig. 1b). The biomass of ANME organisms appears to be accompanied by a large value of e, as would be associated with the oxidation and effective ‘‘fixation’’ of the C1 substrate, CH4, to an enzymatic cofactor (carrier molecule). Although the methane-thiol hypothesis has not yet been confirmed mechanistically, the first isotopic measurements of lipids of ANME Archaea immediately suggested large values of eCH4-biomass (Hinrichs et al., 1999); and numerous followup measurements of lipid values of d13C < 100‰ can only be consistent with the participation of CH4 or other methyl groups as the anabolic substrate(s) for these Archaea. Kerogen in 2.5Ga rocks from the Archean Era, as well as lipids of Archaea from modern and ancient sediments undergoing anaerobic oxidation of CH4, all exhibit similarly depleted values of d13C. This suggests anaerobic oxidation of methane is a quantitatively important process globally. It likely has influenced the sedimentary (and petroleum) record over wide time and spatial scales. A related process is the formation of CH4 by methylotrophic methanogens. Fractionation associated with methylotrophy is summarized in Summons et al. (1998). Notably, the range of fractionation of biomass relative to the methyl group of a likely natural substrate,

151

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trimethylamine, is similar to the fractionation noted for anaerobic oxidation of CH4. Depending on the source of methylated substrates, the biomass of Archaea acting as methylotrophic methanogens in sedimentary communities may not be distinguishable under some circumstances from biomass of ANME species.

3.2

Heterotrophy and Fermentation

3.2.1

Glycolysis

For heterotrophs, the carbon isotopic composition of an organism broadly reflects its carbon source. The d13C values of heterotrophic microbes as well as macrofauna appear to be united by the principle ‘‘you are what you eat, plus 1‰’’ (DeNiro & Epstein, 1978). This rule of thumb is based on the role of the central glycolytic pathway in supplying carbon for intracellular processes (> Fig. 1b). Substrates used by heterotrophs retain their pre-formed carbon isotopic signatures as they are utilized for production of new biomass. The isotopic enrichment of the consumer (þ 1‰) appears to be caused by a weak fractionation associated with the respiratory loss of CO2, i.e., the reverse of the rTCA pathway decarboxylation reactions. In spite of large potential enzymatic fractionations associated with decarboxylation reactions (> Table 1), the fractionation of these enzymes in practice is quite small, with respired CO2 (the product) slightly depleted in 13C relative to the carbon retained by the organism. This rule appears to hold as true for microbial heterotrophs (Blair et al., 1985) as it does for eukaryotes, confirming that the ‘unity of biochemistry’ applies to isotopes and extends throughout life’s Domains. Heterotrophic microbial communities will produce biomass that reflects the preformed 13C content of the organic matter being consumed, and the rule applies irrespective of the electron acceptor utilized (e.g., O2, SO42).

3.2.2

Oxidation of Hydrocarbons

Microbial growth on liquid and gaseous hydrocarbons also conforms to the model above, with the added requirement of additional oxidative steps required to convert subsrates of CxHy to CxHyOz. Typically these pathways are divided into two types of categories: aerobic vs. anaerobic oxidations; and acyl vs. aryl hydrocarbon substrates (> Fig. 1b). Both distinctions highlight the different oxidation states of each of the boundary conditions and the limitations offered by those electron acceptors and donors. The aerobic oxidation of hydrocarbons is mediated by diverse monooxygenases that are similar to pMMO and sMMO in their use of O2 as the electron acceptor. Work on the isotopic discrimination of these enzymes often focuses on the uptake of substrate, rather than on the cells themselves. Fractionation is determined by measuring the time-evolving values of d13C of the residual hydrocarbon, and generally small effects (e <  5‰) are observed (e.g., Morasch et al., 2001). Little is known about the biomass produced from these pathways, but when growing on compounds  C3, the 13C content of biomass should not be greatly different from the starting carbon isotopic composition of the substrate. Similar considerations apply to the anaerobic oxidation of small hydrocarbons. Diverse sedimentary microbes recently were shown to oxidize propane and butane via a proposed initial carboxylation as the activation step (Kniemeyer et al., 2007). Fractionation, measured as

Pathways of Carbon Assimilation and Their Impact on Organic Matter Values d13C

9

discrimination against 13C in the residual substrate, again was generally small (e <  5‰) and scaled with chain length of the substrate (less fractionation at longer chain lengths). Values of d13C of the biomass samples themselves were not measured.

3.2.3

Aceticlastic Methanogenesis and Acetotrophy

Methane produced by aceticlastic methanogenesis is fractionated by 21‰ relative to the methyl group of the acetate from which it is produced (Gelwicks et al., 1994). This implies that knowledge of the pre-formed 13C content of the CH3-group is critical. Uncertainty remains about what fraction of acetigenic methane in natural systems is formed from fermented original organic matter (in which CH3-COOH should reflect the pattern of 13C ordering observed in primary biomass (Monson and Hayes, 1982), most of which usually is formed by the CBB pathway), and how much acetigenic methane derives from the acetate of homoacetogens (in which carbon in CH3-COOH is isotopically homogeneous). The effects above apply to the fractionations between substrate CH3-COOH and CO2 and CH4 products. The mechanism to form biomass in aceticlastic methanogens and acetotrophs is irrespective of the terminal fate of acetate (disporportionation to CO2 and CH4 in the methanogen, or production of two moles of CO2 in the acetate oxidizer). By shuttling a fraction of the substrate to pyruvate via pyruvate synthase (> Fig. 1b), the cell supplies its major intracellular biosynthetic needs. Isotopic values for acetate oxidizers, therefore, should not be governed by fractionation associated with CODH, as the branch point for incorporation of CH3-COOH ! Acetyl-CoA ! pyruvate occurs upstream of CODH (> Fig. 1b). In accordance with this, the biomass of aceticlastic methanogens, and presumably acetotrophs, exhibits minimal fractionation (Londry et al., 2008; > Table 1).

4

Research Needs

Numerous questions raised above require more work to achieve a thorough interpretation of the 13C content of sedimentary organic matter and petroleum. To understand the long-term preservation of organic matter in kerogens and bitumens, the relative fractional contribution of each pathway discussed above must be decoupled, a task that faces nearly infinite challenges. An additional complicating factor for the interpretation of bitumens is the extent to which lipids themselves are fractionated isotopically relative to the total biomass of their original source. Intracellular fractionations result from the diversion of metabolic intermediates such as pyruvate and acetate, which are necessary for the biosynthesis of lipids, into other pathways such as the citric acid cycle and/or for the biosynthesis of amino acids. A detailed mathematical treatment of the isotopic consequences of such branched pathways is given in Hayes (2001), but in general, the fractionation between biomass and lipids (ebiomass-lipid) results in a more negative value of d13C for the lipid. This is true for organisms expressing the Calvin cycle, but it is not necessarily true for species that synthesize acetate directly or for those that have alternative biosynthetic routes to acetate (e.g., van der Meer et al., 2001). Organisms that have unusual metabolisms have not yet been studied thoroughly, and the range of variability in expression of ebiomass-lipid needs further exploration. Finally, to date, little work has been done on organisms for which C1 substrates other than CO2 and CH4 may be important. This includes all instances of methyl carrier molecules as mentioned above, as well as organisms for

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which formaldehyde or formate (e.g., Dolfing et al., 2008; Jahnke et al., 2001), and/or carbon monoxide (King and Weber, 2007), may be significant sources of cellular carbon.

Acknowledgments This review paraphrases many of the contributions of J.M. Hayes, to whom the author is greatly indebted. Support was provided by the David & Lucille Packard Foundation and NSF-EAR.

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10 Global Relations Between the Redox Cycles of Carbon, Iron, and Sulfur W. E. Krumbein1 . A. Gorbushina2 Geomicrobiology, ICBM, Carl von Ossietzky University Oldenburg, Oldenburg, Germany [email protected] 2 LBMPS, De´partement de Biologie Ve´ge´tale, Gene`ve, Suisse

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1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 158 2 Global Biogeochemistry of Carbon, Sulfur, and Iron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 161 3 Hydrocarbons, Carbohydrates, Limestone, Gypsum, and Pyrite Deposits . . . . . . . . . . 162 4 Global Balance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_10, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Solar energy has been transformed into useful redox differences or disequilibria within the Earth’s crust since the onset of anoxygenic and oxygenic photosynthesis in the Precambrian. Inorganic oxidized carbon is transformed into reduced carbon compounds by capturing and storing solar energy. During this process, many different organic compounds are formed including carbohydrates, proteins, hydrocarbons, and various other complex organic metabolic products and their diagenetic polymerization products (melanin, humic substances, petroleum, coal, and kerogen). Many of these solar energy-enriched compounds, however, are oxidized immediately or during the diagenetic transformation of sediments. The oxidation agents are oxygen, sulfate, iron, and other oxidized compounds, which in turn are partly enriched with the original solar energy. On a global biogeochemical scale, however, sulfur and iron are the most important elements. Geological evidence shows that biogeochemical cycles tend to yield stable ratios between the most oxidized forms of carbon (carbon dioxide and calcium/magnesium carbonate) and the most reduced forms (diamond, coal, methane, and hydrocarbons). Throughout the Earth’s history and evolution, this equilibrium ratio is around 1:4, maximally 1:5. When too much carbon is stored in the crust in the form of reduced compounds or vice versa, climatic and biogeomorphogenetic consequences upset the equilibrium. The biosphere reacts in a way to return to the optimal ratio. Excellent examples for this fluctuating equilibrium are the Carboniferous (too much organic carbon stored), the Permian (too little organic carbon stored), and the Tertiary with a generally equivalent production of hydrocarbons and carbonates. At present, we are in a period in which there is a global biogeochemical need to oxidize reduced carbon compounds as fast as possible in order to avoid even more dramatic global climate shifts. The highly evolved human genome seems to be the tool for this shift. Enormous amounts of reduced carbon are turned into the oxidized form as carbon dioxide, which by various biogeochemical pathways is quickly transformed into carbonate, another oxidized form of carbon that can be stabilized and stored in the sedimentary record. Fast recycling of excessively stored solar energy may enable the survival of a global biosphere under highly stressed conditions.

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Introduction

Since the establishment of life on the Earth, biologically mediated processes gained importance for the evolution of the Earth’s crust and upper Mantle. Recently, Falkowski et al. (2008) stated: ‘‘Six major elements – H, C, N, O, S, and P – constitute the major building blocks for all biological macromolecules. The biological fluxes of the first five of these elements are driven largely by microbially catalyzed, thermodynamically constrained redox reactions. These involve two coupled half-cells, leading to a linked system of elemental cycles. On geological timescales, resupply of C, S, and P is dependent on tectonics, especially volcanism and rock weathering. However, reversible metabolic pathways in biogeochemical cycles are not necessarily directly related, and sometimes are catalyzed by diverse, multispecies microbial interactions.’’ Several other important elements, however, are incorporated into these biogeochemical cycles. These are Ca, Mg, Fe, Mn, as well as Si in a complex biologically controlled interactive cycling of redox reactions. > Figure 1 in the communication of Falkowski et al. (2008) clearly outlines the metabolic and energetic relationship between the six major building stones of living matter and the supporting elements. Biological chemosynthesis, anoxygenic and oxygenic photosynthesis, respiration, and disproportioning (fermentation) started and continued to rule the redox status of some

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. Figure 1 In subaquatic and subaerial biological systems (including plankton, benthic biofilms, rock biofilms, and symbiotic systems such as lichen, grass, and forests), enormous amounts of solar energy are captured in the form of reduced carbon and sulfur compounds stored in the Earth crust. Parts of these high-energy carbon compounds are transformed by anaerobic respiration (sulfate reduction) into energy-rich iron and other metal sulfides. Biochemical pathways may, however, release some of this geological energy pool and create carbonates, sulfates, and metal oxides poor in chemical energy. Oxygen-rich reduced carbon compounds represent the center of this biological exchange system of atom migration. The buffering reservoir is represented by oxygen-depleted reduced carbon compounds.

major elements of the global geochemical cycles, the most abundant and most important of which are silica and water. The relationship between free oxygen, water, quartz, and silicates is not very well understood. The cycles of carbon, sulfur, and iron have been more thoroughly studied. The role of a mediator or catalyst is played by phosphorus, a rare element always kept in the fully oxidized status and playing the role of a trigger and controller element, which is cycled at enormous speed without ever changing valence. The organic compounds AMP, ADP, and ATP shift phosphorus through biological processes quickly without ever changing its status from oxidized to reduced. The enormous importance of phosphorus can only be matched by the importance of genomes of biological systems controlling the direction and speed of redox processes of the much more abundant elements, carbon, iron, sulfur, calcium (aragonite, calcite), and magnesium (dolomite). The redox status of all elements mentioned is almost exclusively regulated by the capture of solar energy, as postulated at first by Robert Mayer, one of the founders of thermodynamics (Mayer, 1845). He obviously was the first to realize that the atmosphere constantly rejects huge amounts of solar energy in order to avoid overheating. Simultaneously, however, solar energy is captured in biologically controlled amounts to create useful chemical differences or energetic disequilibria, i.e., potential of directed work. Derived from these introductory remarks, we may now envisage a scenario of interaction between reduced and oxidized carbon, iron, and sulfur that includes the most important cations, calcium and magnesium. Phosphorus as a catalyzer was already mentioned as well as the still enigmatic positioning of silica and water

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. Figure 2 (a) The young Earth exhibits an oxygen-free atmosphere. Heavy cloud cover and volcanic exhalations through thin crust layers reduce solar heat input to a minimum. Primordial radioactivity and accretion heat reach the surface and keep surface temperatures high. With decreasing internal energy supply, increasing amounts of liquid water and solar luminosity ( Fig. 1 are controlled by solar energy input via photophosphorylation and subsequent metabolic cycles of living matter. For example, an increase in the amount of sulfur occurring in the oxidized state as calcium sulfate (anhydrite or gypsum) is accompanied by an increase in the amount of iron occurring in the oxidized state (goethite or hematite). Typical minerals of this interaction between carbon, iron, sulfur, and microbial energy and electron needs are schwertmannite (ideal formula Fe8O8(OH)6(SO4)) and jarosite ((K,NH4,H3O)Fe3(SO4)2(OH)6), which are cycled by sulfur-oxidizing and sulfur-reducing bacteria. When sulfur turns into the reduced state, also the iron oxides, goethite and hematite, will simultaneously turn into iron and other metal sulfides (e.g., pyrite) (Lovely et al., 2004; Wang et al., 2007). Sulfate is reduced and serves as an electron acceptor liberating carbon dioxide and calcium. These will eventually form calcium and/or magnesium carbonates. Herein direct geological evidence is outlined for the enormous importance of the intimate relationships among carbon, sulfur, and iron. These need to be analyzed on a global scale not only for present-day conditions, but also historically in order to clarify the importance of mass balances for climate and habitability of the Earth in the past and the present. In order to achieve this, one must look at (1) reservoirs (atmosphere, ocean, sediment, crust, and mantle), and (2) residence times and fluxes from a daily scale to geological timescales (Walker, 1993). One must obtain quantitative data on the amounts of energy stored and released through geological time from the reduction and oxidation of iron and sulfur, as related to photosynthesis products, and of geochemical data gained from the analysis of sedimentary systems through geological time (Kasting and Walker, 1993; Krumbein and Schellnhuber, 1992). Further thermodynamics of complex relationships in living ecosystems need to be considered in order to understand how geological timescales are interlinked

form of hydrocarbons, kerogen, and sulfides through partial organic matter oxidation via nitrate, iron, manganese, and sulfate respiration. The resulting sulfides remain as energized reservoirs for future geophysiological activity. Heat storage from above initiates top-down tectonics while internal cooling creates thicker and more stable crustal elements ready for recycling. Disequilibria of these processes are reflected in warm/wet/reduced geological periods (Carboniferous) with excessive reduced carbon production (coal, oil, etc.) and sulfide ores and cold/dry/oxidized periods (Rotliegend/Permian) with excessive production of metal oxides, gypsum, and limestone. The primary and secondary productivity-related ratios of reduced to oxidized carbon, thus, may have oscillated throughout geological periods of time and may have also induced or accelerated ice age periods (near Snowball worlds).

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with biological dynamics (Joergensen and Svirezhev, 2004). ‘‘Newton’s Laws,’’ Maxwells law of electricity, Quantum theory, Einstein’s Relativity theories, thermodynamic laws, and laws of radiation need to be incorporated into the analysis of biogeochemical cycles on a geological timescale once they are embedded in biological time. All of them need to be time embedded again into a number of laws covering physical relationships between elements and life as a driving force (Krumbein, 1983; Westbroek, 1991; Gerdes and Krumbein, 1987). From these multiple approaches and assumptions, one may, ultimately, come to the conclusion that not only the intimate relationship between the cycles of carbon, sulfur, and iron is under the control of living systems, but also the dynamics of the Earth itself (Anderson, 1984; Krumbein, 2008). As postulated by Vernadsky (1929), even the migration and close association of all atoms available in the Earth’s crust and upper mantle are accelerated, retarded, or organized in space and time by the force of living matter.

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Hydrocarbons, Carbohydrates, Limestone, Gypsum, and Pyrite Deposits

How do the cycles of carbon, sulfur, and iron really interact? According to the view of some biogeoscientists (Anderson, 1984; Krumbein, 2008; Rosing et al., 2006), the amount of energy stored in rock deposits is the clue to this question. Photophosphorylation annually captures a certain percentage of solar energy, and transforms it into energy-rich reduced carbon compounds. Only 1% of this production of energy-rich organic compounds is not recycled by annual respiratory activity of the total biosphere and is stored in litter, peat, soil, and sediment organic matter (coal, petroleum, methane, kerogen). This, however, may be sufficient to keep the crust and upper mantle under the control of living matter. This represents only 0.5% of the total mass of the Earth. This uppermost 30–80 km thick and motile layer of rock and magma, however, is most important for biogeochemical and geological cycles and guarantees the survival of a biosphere on the Earth. As Anderson (1984) stated: "

‘‘It has often been suggested that life established on earth because of a coincidence between the narrow temperature interval over which water is liquid and the temperature extremes that actually occur on earth. The earth apparently is also exceptional in having plate tectonics (or platonics cf. Krumbein, 2008). If the carbon in the atmosphere of Venus could turn into limestone, the surface temperatures and those of the upper mantle would drop. The basalt eclogite phase change would migrate to shallow depths, causing the lower part of the crust to become unstable. Thus there is the interesting possibility that plate tectonics may exist on earth because limestone generating life established itself on this planet.’’

On this global and the Earth historical scale, several important facts have apparently been overlooked by modern ecologists and global climate modelers analyzing energy reservoirs, fossil carbon burning, and global climate consequences. These are 1. More than 98% of presently living biomass is represented by microorganisms (bacteria, algae, fungi). 2. More than 99% of the history of life on earth has been organized and controlled by microorganisms (even during the anoxic period???). 3. Cycling (redox reactions using solar energy) has been achieved exclusively by bacterial chemosynthesis and photosynthesis aided very early (beginning in the Precambrian) by

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fungal mineralization of reduced carbon compounds and fungal support of photosynthesis through symbiotic cooperation (lichen, plants, trees, forests). 4. Still today in the electric era, more than 99% of all energy transfers and redox reactions are performed by microbes. Examples are sedimentary biofilm, rock biofilm, and continental shelf transformations of oil and methane seepages at global scales exceeding the total energy cycling and waste by human technical manifestations. 5. Microbial life and metabolic activities continue to cycle energy through photosynthetic and respiratory redox reactions keeping the amount of solar energy stored in the Earth’s crust relatively constant. 6. Climate and habitability are regulated by biogeochemical and biogeomorphogenetic processes in which the rapid wear-down of subaerial continental surfaces play the same key role as the production of reduced carbon compounds and in turn of oxidized carbonate rocks. In order to finally underscore these statements, it seems to be appropriate to remind ourselves of the orders of magnitude of these microbial reactions.

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Global Balance

Kempe (1979) and the CRC handbook of Chemistry and Physics through 50 continuous editions give the following amounts for carbon in the Earth’s crust. They assume 60  1021 g carbonate carbon balanced by 12  1021 g organic carbon compounds with a ratio of 5:1–4:1 throughout the geological history. A problem is presented by the different turnover times between 12 years in the biosphere and about 300 million years in the lithosphere. The energetic relations and biogeochemical cycles may be of geological and climate relevance on all timescales. If we assume an average value of 3,000 kJ free energy for an average organic carbon compound (sugar), we will arrive at values between 15.0  1023 and 3.6  1024 kJ energy fixed in organic carbon compounds and reduced (sulfide) metal compounds. The annual solar energy capture rate (top-down) in organic carbon compounds and energy-rich sulfides derived from sulfate respiration matches approximately the average annual heat flow of the crust (bottom-up). The most moderate assumption derived from these figures is that solar energy powered organic carbon and sulfide energy budgets and flows in the crust may be of the same order of magnitude as the magma-derived flows powered by the upper mantle of the Earth. In > Figs. 2a and b, an attempt is made to compare a primordial lifeless Earth with little or no mobility of crustal elements with the Earth powered at least partially by captured solar energy with consequent limestone production and mobile and flexible crustal elements, which, in turn, enable nutrient replenishment and climate regulation. Not unlike the rowboat propelled on a thin water layer by human muscle power, flexible plates in the game of continental movements (horizontal and vertical) may be translocated using energies derived from solar power stored in organic carbon compounds and sulfide ores. Hereby, a scenario of a mobile crust and the turnover of elements necessary for living matter is created that matches the early writings of Herder, Kant, and A. von Humboldt (Krumbein and Schellnhuber, 1992; Krumbein, 1996) and the most recent approach of Anderson (2007). Thus, in a global outlook at the cycles and meeting points of carbon and sulfur, we may need to step forward from biogeochemistry and biogeochemical cycles to geophysiology and a planetary metabolism under the control of living matter and biogenic migration of atoms (Vernadsky, 1929).

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An additional view may also be that, in the production and conservation of organic carbon compounds, the sulfur cycle also plays an important role. Pickling of vegetable (cucumber) is an old tradition of making preserves. Pickling again is typical for the production and conservation of hydrocarbons in hypersaline environments. Within the same environments, huge amounts of gypsum and calcium carbonate (limestone) are formed. These in turn are kept sealed by clay minerals produced by climate-derived bioerosion and rock destruction (Gorbushina, 2007). This leads to the accumulation of porous storage rocks capped by impermeable clay layers in the vicinity of oil production sites. Hydrocarbons migrate into these rocks and are covered by huge salt deposits (NaCl, MgCl) and topped by sheet flood-derived clay and silt deposits. Hereby, large differences in the specific weight of rock materials are created facilitating local and global plate or continent movements. Hydrocarbons may, thus, (in connection with gypsum and limestone deposits) also serve as a lubrificant not unlike the oil used to keep mobile parts of machines (cars) in constant action. In order to summarize some of the data and thoughts presented so far instead of a discussion or a summary, we put together the step-by-step scenario of carbon, sulfur, iron, and water and silica interactions in a concluding table. The cycles of carbon and sulfur, thus, are intimately

. Table 1 Important geophysiological operations of the bioplanet (Modified from Krumbein and Schellnhuber, 1992) 1. Solar-powered anoxygenic and oxygenic photosynthesis creates energy-rich carbon compounds (including 99% kerogen and 1% petroleum and methane) 2. Equilibrated simultaneous production of carbonates (limestone, dolomite) 3. Respiratory and disproportionating (fermentative) action of living matter recycles 99% and deviates some of the energy into energy-rich sulfides. About 1% of the annual production is stored for geological periods of time (carbon and sulfur are at a ratio of 10:1) 4. Living matter controls weathering and solution (oxidation) rates and hereby controls geomorphology via biogeomorphogenesis (Naylor and Viles, 2002) 5. Total control of albedo, i.e., regulation of backscattering of solar radiation by keeping atmospheric composition at relatively constant levels via living matter biogeochemistry 6. Absolute control of phosphorus compounds and their cycling speed by bioenergetics 7. Partial (?) control of the chemical and geomechanical silica and water cycles 8. Speed of biological migration of atoms adapted to external energy stress (changing solar radiation rate) 9. Accelerated mass transfer (horizontal and vertical) via macroorganism evolution 10. Total control of inflow, capture, storage, and release of solar energy via hydrocarbon to carbonate and metal sulfide to metal oxide ratio balancing, powered by photosynthesis, respiration, disproportioning (fermentation), passive (migration), and active transport (swimming, running, flying) of atoms 11. Control on rock densities and atom distribution in atmosphere, hydrosphere, lithosphere, and mantle 12. Bringing carbon, sulfur, and iron cycles under control of living matter (also silica and water?); work needs to be done on this

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linked to those of calcium, iron, phosphorus, and other essential elements at local and planetary scales as well as on short-term and extended (geological) timescales (> Table 1).

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Research Needs

Within the frame of clearly established reversible metabolic pathways connecting the redox reactions of carbon, sulfur, iron, and calcium on a global scale, some hypothetical assumptions need to be further elucidated. Most important: Does annual solar energy capture and deposition in sedimentary rocks via reduced carbon and sulfide compounds really reach the same order of magnitude as the geothermal heat flow registered, e.g., in deep mines and by geothermal measurements? Moreover, it seems that the cycles of silica and water are connected to the flow of solar energy. Water is transformed into free oxygen and reducing power and reproduced by respiration. Any reduced carbon compound stored in rock material will not

. Figure 3 The solar energy pump charges huge amounts of quanta into reduced organic carbon compounds. Some of the energy is transmitted by sulfate reduction into energy-rich sulfides. Organic carbon and sulfide ores may be oxidized with oxygen or by metabolic processes and release the solar energy captured million years before.

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immediately recycle into water. The amounts of water cycling through the biota, however, may be negligible. Huge amounts of silica are cycled, and again, the biological input into the silica cycle seems to be negligible. On the other hand, the cycles of silica and water seem to have little connection yet to those of carbon, iron, and sulfur in biogeochemical considerations. The deviations of element distribution considering the universe, the whole Earth, and its crust are considerable. If we compare the whole Earth and its crust, only we see major disparities in the following elements (whole earth/crust in percentage). Iron 35/5; oxygen 30/46; silicon 15/27; magnesium 13/2; sulfur 1.9/0.5; calcium 1.1/3.6; and carbon unknown/ 0.32. Phosphorus and manganese lie in the order of 0.1%. The biologically motivated migration, accumulation, and residence times of all these elements in the Earth’s crust are not yet fully analyzed or understood. Seemingly, however, more and more exact data are urgently needed. Long- and short-term consequences of biological cycling of carbon, sulfur, and water are also depicted in a partially humoristic way in > Figs. 3–6. Most of the data indicated in the Handbook of Chemistry and Physics have not been changed since almost 50 years. Most of them have been collected in Institutes of the former Soviet Union Academy

. Figure 4 The relations between external energy, original internal energy, and variable amounts of solar energy stored in sediments and crustal rocks. The amount of stored solar energy varies in different geological eras and formations.

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. Figure 5 A kind of caricature of behavior of a bioplanet on geological scale. Earth is warming, when methane, oil, coal, and kerogen are oxidized and reduced energy-rich carbon compounds are transferred by metabolic oxidation or burning (fire), into the oxidized forms of carbon dioxide and earth alkali carbonates (see > Fig. 1). In turn, Earth is cooling, when large amounts of carbon dioxide and calcium carbonate are transformed into reduced energy-rich carbon compounds. The processes seem to be self-regulatory over geological periods of time. The acceleration rate by human activity presently is difficult to assess.

of Science with thousands of mineralogists and geochemists analyzing rocks of different geological periods. We urgently need not only global climate modelers, but also global crustal evolution and dynamic analysts. Fortunately, a new generation of biogeoscientists combining microbiology, geochemistry, and thermodynamics has emerged in the past 30 years since the first Geomicrobiology Chair worldwide was established at Oldenburg University in 1979.

Acknowledgments The authors acknowledge support by a Feodor Lynen scholarship of Alexander von Humboldt Foundation to AAG and a symposium grant of DFG to WEK. We further gratefully acknowledge Frances Westall, Orle´ans (France), and James MacAllister, Amherst, Massachusetts (USA) for cross-reading and valuable suggestions.

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. Figure 6 Fine tuning of biological impacts has been demonstrated by physicists. The rotational speed of Earth increases on a measurable scale by pumping water several tens of metres up into the foliage of trees and forests in (especially northern) spring and summer and decreases in turn, when the foliage is falling down in fall and winter. Although marginal, the effect is measurable and is an excellent example of global geophysiology on a smaller time scale than production and consumption of reduced carbon compounds over geological periods of time.

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Falkowski PG, Fenchel T, Delong EF (2008) The microbial engines that drive earth’s biogeochemical cycles. Science 243: 1034–1039. Friedman GM, Krumbein WE (1985) Hypersaline ecosystems. The Gavish Sabkha. Berlin: Springer, 484 p.

Global Relations Between the Redox Cycles of Carbon, Iron, and Sulfur Gerdes G, Krumbein WE (1987) Biolaminated Deposits. Berlin: Springer, 183 p. Gorbushina AA (2007) Life on the Rocks (Mini-Review). Environmental Microbiology 9: 1613–1631. Joergensen SE, Svirezhev YM (2004) Towards a Thermodynamic Theory for Ecological Systems. Amsterdam: Elsevier, 366 p. Kasting JFD, Walker JCG (1993) Long-term effects of fossil-fuel-burning and deforestation on levels of atmospheric CO2. pp. 151–165. In: Oremland RS (ed.). Biogeochemistry of Global Change. London: Chapman & Hall, 879 p. Kempe S (1979) Carbon in the rock cycle. In: Bolin B, Degens ET, Kempe S, Ketner P (eds.). The Global Carbon Cycle Scope, 13. New York: Wiley, 343 p. Krumbein WE (1983) Microbial Geochemistry. Oxford: Blackwell, 333 p. Krumbein WE (1996) Geophysiology and parahistology of the interactions of organisms with the environment. Marine Ecology 17: 1–21. Krumbein WE (2008) Biogenerated rock structures. Space Sci Rev 135: 81–94. Krumbein WE, Schellnhuber HJ (1992) Geophysiology of mineral deposits – a model for a biological driving force of global changes through Earth history. Terra Nova 4(3): 351–363.

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Lovley DR, Holmes DE, Nevin KP (2004) Dissimilatory Fe(III) and Mn(IV) reduction. Adv Microb Physiol 41: 221–286. Mayer JRv (1845) Die organische Bewegung in ihrem Zusammenhange mit dem Stoffwechsel. Ein Beitrag zur Naturkunde. Heilbronn: Drechsler, 52 p. Naylor LA, Viles HA, and Carter NEA (2002) Biogeomorphology revisited: Looking towards the future. Geomorphology 47: 3–14. Rosing M, Bird DK, Sleep NH, Glassley W, Albarede F (2006) The rise of continents – an essay on the geologic consequences of photosynthesis. PALAEO 232: 199–213. Vernadsky VI (1929) The Biosphere. New York: Springer/ Kopernikus, 192 p. Walker JCG (1993) Biogeochemical cycles of carbon on a hierarchy of time scales. pp. 3–28. In: Oremland RS (ed.). Biogeochemistry of Global Change. London: Chapman & Hall, 879 p. Westbroek P (1991) Life as a Geological Force. New York: Norton & Co., 240 p. Wang H, Bigham JM, Tuovinen OH (2007) Synthesis and properties of ammoniojarosites prepared with ironoxidizing acidophilic microorganisms at 22 to 65 C. Geochim Cosmochim Acta 71: 155–164.

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11 History of Life from the Hydrocarbon Fossil Record C. C. Walters1 . K. E. Peters2 . J. M. Moldowan3 1 ExxonMobil Corporate Strategic Research, Annandale, NJ, USA [email protected] 2 U.S. Geological Survey, Menlo Park, CA, USA [email protected] 3 Department of Geological & Environmental Sciences, Stanford University, Stanford, CA, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172 2 Methanogenic Archaea in the Archean . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172 3 Cyanobacteria and the Great Oxidation Event . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 173 4 Early Eukarya and Steranes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 174 5 Biomarkers of the Great Dying . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 175 6 Diatoms, 23,24-Dimethylcholestanes and HBIs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 176 7 Higher Land Plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 177 8 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 179

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_11, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Certain lipids and biopolymers retain their original structure through sedimentary diagenesis and catagenesis that they can be assigned to a specific biological origin. These ‘‘taxon-specific biomarkers’’ (TSBs) can serve as chemical fossils that trace the evolution of life. TSBs in Early Precambrian rocks reveal the early evolution of archaea, cyanobacteria, and eukarya and the development of atmospheric free oxygen. Steroidal TSBs document the changing nature of marine phytoplankton from Neoproterozoic organic-walled acritarchs to the predominance of present day diatoms and terpanoid TSRs reveal the evolution of higher land plants. TSBs, used in conjunction with isotopic analysis, can identify the taxa of enigmatic fossils, provide important clues to the causes of mass extinctions, and describe the global changes in biotic diversity and Earth’s conditions as the biosphere recovers. Biomarkers record the evolutionary history of life on Earth and perhaps, other planets.

1

Introduction

Most of life’s chemical components are unstable when deposited in sediments. DNA degrades very rapidly and is restricted to fossils younger than 50 thousand years old preserved in permafrost. Proteins are more stable, but typically lose their diagnostic sequences within 102–105 years. Collagen proteins are stabilized in fossil bones (Collins et al., 2000) and fragments have been successfully sequenced from a 160,000- to 600,000-year-old fossilized mastodon and possibly an exceptionally well preserved, 68 million-year old T. Rex femur (Asara et al., 2007). Many lipids are stable and can survive in sediments for a much longer time (Volkman, 2006). For example, alkenones, which are produced by Prymnesiophyceae phytoplankton, are highly resistant and have been found in sediments as old as 120 Million years (Brassell et al., 2004). Most lipids do not remain intact during burial and lithification of sediments. When exposed to higher temperatures, these compounds undergo chemical and microbial alteration involving the loss of functional groups and hydrogenation, cleavage, cross-linking, condensation, and aromatization reactions. However, certain lipids and biopolymers retain enough of their original molecular structure to assign a specific biologic origin (Peters et al., 2005). These hydrocarbons, termed biomarkers by the geochemical community, can endure exposure to higher temperatures and survive for billions of years. Some ‘‘taxonspecific biomarkers’’ (TSB; Moldowan and Jacobson, 2000) have structures that are specific to a taxonomic group and can serve as chemical fossils that trace the evolution of life. A comprehensive review of biomarkers and their evolution is beyond the scope of this paper and the reader is directed to several recent publications that more broadly cover these topics (Brocks and Pearson, 2005; Brocks and Summons, 2003; Moldowan and Jacobson, 2000; Peters et al., 2005; Summons and Walter, 1990 ). Instead, we will examine several recent examples that illustrate the use and limitations of biomarkers as chemical fossils.

2

Methanogenic Archaea in the Archean

Ribosonal sequencing suggests that the three kingdoms of life, archaea, eubacteria and eukarya, split from a common ancestor 4000 Ma (Ma = 106 years) (Hedges, 2002; Sheridan et al., 2003). Alternative theories suggest that archaea and eukarya evolved much later 900 Ma (Cavalier-Smith, 2002, 2006). The chemical record favors an early divergence. Tailto-tail linked C20 (2,6,11,15-tetramethylhexadecane = crocetane) and C25 (2,6,10,15,19pentamethylicosane = PMI) isoprenoid hydrocarbons are diagnostic markers for methanogenic

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archaea or anaerobic methanotrophic consortia and one or both of these compounds are reported to occur in shale extracts from the 750 Ma Chuar Group (Summons et al., 1988a) and the 1690 Ma Barney Creek Formation (Greenwood and Summons, 2003; Summons et al., 1988b). Extracts from 2707 to 2685 Ma metasedimentary rocks from Timmins, Ontario, Canada contain cyclic and acyclic biphytanes and C36-C39 derivatives (Ventura et al., 2007). These compounds are likely to be syngenetic as similar compounds were released by catalytic hydrogenation of the extracted rock. Archaeal biomarkers are supported by light stable carbon isotope ratios (< 55‰) measured for Archean kerogens that are presumably derived from anaerobic methantrophs (Eigenbrode and Freeman, 2006). These findings also agree well with some estimates of when methanogens diverge within the Euryarchaeota (Sheridan et al., 2003). Isotopically light methane in 3500 Ma fluid inclusions from the Pilbara craton may be evidence for a very early divergence (Ueno et al., 2006), but it is unclear if this methane is biogenic (Sherwood Lollar et al., 2006).

3

Cyanobacteria and the Great Oxidation Event

For over two billion years, the earth’s atmosphere contained little or no oxygen (Canfield, 2005). Between 2450 and 2320 Ma, the oxygen level rapidly increased to about one percent (Farquhar et al., 2007; Holland, 2006). The transition to an oxic atmosphere, termed the Great Oxidation Event, resulted because free oxygen produced by cyanobacteria outpaced its sequestration in mineral oxides (Kump and Barley, 2007). When cyanobacteria evolved, however, remains uncertain. RNA phylogenetic analysis suggests that major diversification of eubacteria, including the cyanobacteria, took place 2600  300 Ga (Hedges, 2002; Sheridan et al., 2003). The Great Oxidation Event could then have occurred shortly after cyanobacteria evolved, rapidly producing free oxygen that destroyed greenhouse methane and precipitated a snowball Earth (Kopp et al., 2005). Microfossils and stromatolites, however, suggest that cyanobacteria existed long before the Great Oxidation Event (Allwood et al., 2006; Schopf, 2006). Although claims for cyanobacterial fossils dating back 3500 Ma are disputed (Brasier et al., 2006), more convincing evidence exists in rocks 3000–2900 Ma (Altermann and Kazmierczak, 2003; Nisbet et al., 2007; Schopf et al., 2007). The oldest microfossils with definitive cyanobacterial features date no earlier than 2100 Ma (Hofmann, 1976; Tomitani et al., 2006). The discovery of 2-methylhopanes in extracts from 2700 Ma (Brocks et al., 1999) and 2780 Ma (Brocks et al., 2003a, b) Pilbara shales appeared to prove that cyanobacteria evolved significantly prior to the Great Oxidation Event. 2b-methylhopanoids are biosynthesized by cyanobacteria and their corresponding sedimentary 2a-methyhopanes were considered as TSBs for this phylum (Summons et al., 1999). However, the taxonic specificity of 2-methylhopanes must be re-examined because their precursors are now known to be synthesized by strains of Rhodopseudomonas palustris, a purple non-sulfur phototrophic a-proteobacterium (Rashby et al., 2007). DNA-sequencing studies indicate that bacterochlorophylls evolved prior to the chlorophyll in cyanobacteria (Xiong et al., 2000). In the biosynthesis of chlorophyll, aerobes use an enzyme that incorporates molecular oxygen, anaerobes use an enzyme that incorporates oxygen from water and some facultative phototrophs and can utilize either pathway depending on their environment. Therefore, the proto-cyanobacteria must have utilized anaerobic enzymes to synthesize chlorophyll, which in turn, were replaced in cyanobacteria by the aerobic enzymes

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as molecular oxygen became available (Raymond and Blankenship, 2004). R. palustris has a remarkably diverse set of genes supporting four modes of metabolism, which allows it to exist as a photoautotroph or photoheterotroph in the absence of oxygen or as a chemoheterotroph or chemoautotroph when oxygen is present (Larimer et al., 2004). The ability of R. palustris to biosynthize 2-methylhopanoids could be the result of horizontal gene transfer or it could indicate that this extant microbe is related to the ancestor that gave rise to cyanobacteria. This would allow cyanobacteria to evolve after the Great Oxidation Event as suggested in some molecular phylogenetic studies (Tomitani et al., 2006).

4

Early Eukarya and Steranes

The biosynthetic pathways for building hopanoids or sterols represent a major, but not universal, distinction between eubacteria and eukaryotes. Both pathways begin with the synthesis of squalene that is (1) folded into hopanoids or (2) enzymatically linked with molecular oxygen forming epoxide 2,3-oxidosqualene, which is then folded into one of the two protosterols, lanosterol or cycloartenol (Volkman, 2005). Further modifications involve a series of cyclizations and subsequent oxidative demethylations by enzymes requiring additional O2. The folding cyclases are remarkably similar, indicating that it is the insertion of molecular oxygen in squalene that marks the evolutionary divergence (Fischer and Pearson, 2007; Summons et al., 2006a). The oldest known Eukarya microfossils are only 1900 Ma (Javaux 2007; Knoll et al., 2006). As all modern eukarya utilize sterols, the detection of steranes in Archean and Early Proterozoic rocks implies an earlier evolution and the presence of molecular oxygen. An indigenous origin for the steranes in the >2.5 Ma Pilbara craton shales has been questioned (Kopp et al., 2005), so we will limit our discussion to the steranes found in fluid inclusions within 2450 Ma Matinenda Formation quartz (Dutkiewicz et al., 2006; George et al., 2008). The inclusions contain a full range of saturated and aromatic hydrocarbons that resemble produced oil from the Late Precambrian source rocks. In addition to abundant biomarkers that are consistent with cyanobacteria (e.g., 2-methylhopanes, monomethyl alkanes), a diverse suite of steranes occur at lower concentrations. C28 and C29 steranes and the C28 diasteranes are unambiguously indigenous. C27, C30 24-n-propyl-, C30 4a-methyl-, and C26 24-norand 27-nor-cholestanes and diacholestanes also were detected, while dinosteranes and 24-isopropylcholestanes were not. The regular C27, C28, and C29 diasteranes and steranes are not taxon specific beyond being characteristic of eukarya. C30 24-n-propylcholestanes originate from sterols in some chrysophytes (Moldowan et al., 1990) while 24-isopropylcholestanes originate from sponges and are particularly dominant in the Late Vendian-Early Cambrian source rocks (McCaffrey et al., 1994); thus, chrysophytes are implicated as the source of 24-npropylcholestanes in the Matinenda Formation oil inclusions. Collectively, the C30 4a-methylsteranes and C26 norcholestanes tell an interesting story. C30 4a-methylsteranes can originate from multiple sources, but are most commonly associated with dinoflagellates. One specific type, the dinosteranes, are produced almost exclusively by dinoflagellates. The fossil record for dinoflagellates begins in the Triassic, but because few living dinoflagellates produce fossilizable cysts; this record has long been only a suspicion. Organic-walled acritarchs believed to be related to dinoflagellates (Moldowan et al., 1996) date back to the Neoproterozoic (Butterfield and Rainbird, 1998). Here, the mineralized fossil record matches well with the chemical fossil observations as dinosteranes and related aromatic

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biomarkers have a continuous record dating back to the Early Cambrian (Moldowan and Talyzina, 1998) and possibly the Neoproterozoic (Summons et al., 1992; Zhang et al., 2002), but are most abundant in post-Paleozoic sediments. The chemical record for C26 24-norcholestanes and 24-nordiacholestanes also dates back to the Neoproterozoic (Zhang et al., 2002), although concentrations of these compounds in source rocks remain low until the Jurassic and become particularly abundant in the Tertiary (Holba et al., 1998a, b). This temporal pattern and the association of high concentrations of 24-norcholestanes with siliceous sediments containing diatom-specific biomarkers (e.g., highly branched isoprenoids) strongly suggests that these compounds are from diatoms. However, diatoms did not evolve until 100 diatom species found abundant 23,24-dimethylsterols in twenty-one diatoms belonging to six different orders (Rampen et al., 2007b). The appearance of triaromatic 23,24-dimethylcholestanes in the Triassic with increasing abundance through the Cretaceous is consistent with diatom and dinoflagellate sources. Diatoms of the genera Rhizosolenia, Haslea, Navicula, and Pleurosigma are the only organisms known to synthesize highly branched isoprenoids (HBIs). The function of these ‘‘T-branched’’ alkenes is unknown, but two groups of diatoms evolved independent pathways for their synthesis implying they provide a significant advantage (Masse´ et al., 2004). In a study that integrated 18S rRNA phylogenetic analysis with the mineral and chemical fossil record, Sinninghe Damste´ et al. (2004) showed that HBIs were first biosynthesized by the rhizosolenid diatoms. The mineral fossil record for the rhizosolenids dates to 70 Ma, while saturated HBIs are found in oils and marine sediments 2 billion years. Geochim Cosmochim Acta 72: 844–870.

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sediment of the late Proterozoic Walcott Member, Chuar Group, Grand Canyon, Arizona. Geochim Cosmochim Acta 52: 2625–2637. Summons RE, Jahnke LL, Hope JM, Logan GA (1999) 2-Methylhopanoids as biomarkers for cyanobacteial oxygenic photosynthesis. Nature 400: 554–557. Summons RE, Love GD, Hays L, Cao C, Jin Y, Shen SZ, Grice K, Foster CB (2006b) Molecular evidence for prolonged photic zone euxinia at the Meishan and East Greenland sections of the Permian Triassic Boundary. Geochim Cosmochim Acta 70: A625. Summons RE, Powell TG, Boreham CJ (1988b) Petroleum geology and geochemistry of the Middle Proterozoic McArthur Basin, Northern Australia: III. Composition of extractable hydrocarbons. Geochim Cosmochim Acta 52: 1747–1763. Summons RE, Thomas J, Maxwell JR, Boreham CJ. (1992) Secular and environmental constraints on the occurrence of dinosterane in sediments. Geochim Cosmochim Acta 56: 2437–2444. Summons RE, Walter MR (1990) Molecular fossils and microfossils of prokaryotes and protists from Proterozoic sediments. Am J Sci 290-A: 212–244. Taylor DW, Li H, Dahl J, Fago FJ, Zinniker D, Moldowan JM (2006) Biogeochemical evidence for the presence of the angiosperm molecular fossil oleanane in Paleozoic and Mesozoic non-angiospermous fossils. Paleobiol 32: 179–190. Tomitani A, Knoll AH, Cavanaugh CM, Ohno T (2006) The evolutionary diversification of cyanobacteria: Molecular-phylogenetic and paleontological perspectives. Proc Natl Acad Sci 103: 5442–5447. Ueno Y, Yamada K, Yoshida N, Maruyama S, Isozaki Y (2006) Evidence from fluid inclusions for microbial methanogenesis in the early Archaean era. Nature 440: 516–519. Ventura GT, Kenig F, Reddy CM, Schieber J, Frysinger GS, Nelson RK, Dinel E, Gaines RB, Schaeffer P (2007) Molecular evidence of Late Archean archaea and the presence of a subsurface hydrothermal biosphere. Proc Natl Acad Sci 104: 14260–14265. Volkman JK (2005) Sterols and other triterpenoids: source specificity and evolution of biosynthetic pathways. Org Geochem 36: 139–159. Volkman JK (2006) Lipid markers for marine organic matter. In The Handbook of Environmental Chemistry, JK Volkman (ed.). Reactions and processes, Part N, Marine Organic Matter: Biomarkers, Isotopes and DNA. vol. 2. Berlin: Springer, pp. 27–70. Volkman JK, Eglinton G, Corner EDS (1980) Sterols and fatty acids of the marine diatom Biddulphia sinensis. Phytochemistry 19: 1809–1813. Wang C Liu Y Liu H Zhu L, Shi Q (2005) Geochemical significance of the relative enrichment of pristane

History of Life from the Hydrocarbon Fossil Record and the negative excursion of d13CPr across the Permian-Triassic boundary at Meishan, China. Chin Sci Bull 50: 2213–2225. Wang C, Visscher H (2007) Abundance anomalies of aromatic biomarkers in the Permian-Triassic boundary section at Meishan, China – Evidence of end-Permian terrestrial ecosystem collapse. Palaeogeogr Palaeoclimatol Palaeoecol 252: 291–303. Waters ER (2003) Molecular adaptation and the origin of land plants. Mol Phylogenet Evol 29: 456–463. Wellman CH, Osterloff PL, and Mohiuddin U (2003) Fragments of the earliest land plants. Nature 425: 282–285. Withers N (1983) Dinoflagellate sterols. In Marine Natural Products 5. PJ Scheuer (ed.). New York: Academic Press, pp. 87–130. Xie S, Pancost RD, Huang J, Wignall PB, Yu J, Tang X, Chen L, Huang X, Lai X (2007a) Changes in the global carbon cycle occurred as two episodes during the Permian–Triassic crisis. Geology 35: 1083–1086.

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Xie S, Pancost RD, Huang X, Jiao D, Lu L, Huang J, Yang F, Evershed RP (2007b) Molecular and isotopic evidence for episodic environmental change across the Permo/Triassic boundary at Meishan in South China. Glob Planet Change 55: 56–65. Xie S, Pancost RD, Yin H, Wang H, Evershed RP (2005) Two episodes of microbial change coupled with Permo/Triassic faunal mass extinction. Nature 434: 494–497. Xiong J, Fischer WM, Inoue K, Nakahara M, Bauer CE (2000) Molecular evidence for the early evolution of photosynthesis. Science 289: 1724–1730. Yin H, Feng Q, Lai X, Baud A, Tong J (2007) The protracted Permo-Triassic crisis and multi-episode extinction around the Permian-Triassic boundary. Glob Planet Change 55: 1–20. Zhang S, Moldowan JM, Li M, Bian L, Zhang B, Wang F (2002) The abnormal distribution of the molecular fossils in the pre-Cambrian and Cambrian: its biological significance. Science China (Series D) 45: 193–200.

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Part 3

Transfer from the Geosphere to Biosphere

12 Marine Cold Seeps E. Suess Leibniz-Institute for Marine Sciences (IFM-GEOMAR), Kiel, and German Marine Research Consortium (KDM), Berlin, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 188

2 2.1 2.2 2.3

Seeps at Active Plate Margins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Oceanic Plate and Continental Plate Convergence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Oceanic Plate and Oceanic Plate Convergence . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 189 Transform Plate Boundaries . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

3

Seeps at Passive Continental Margins and Shelves . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 191

4

Common Seep Characteristics at Active and Passive Margins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 192 Sedimented Margins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 Formation and Manifestations of the Hydrocarbon-MetazoanMicrobe-Carbonate Association . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 193 Gas Hydrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194 Anoxic Oxidation of Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 194 Authigenic Carbonates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 Biota . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 The Black Sea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 195 Evaporitic Host-Strata: The Gulf of Mexico and The Eastern Mediterranean Sea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196

4.1 4.1.1 4.1.2 4.1.3 4.1.4 4.1.5 4.1.6 4.2

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Deep Fluids from Erosive and Transform Margins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 196

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Research Needs: Significant Issues of Seep Research . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Global Methane Emissions from Seeps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Ancient Seeps and Fossilization of Microbial Structures . . . . . . . . . . . . . . . . . . . . . . . . . . 197 Benthic Microbial Fuel Cell at seeps . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 198

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_12, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Marine seeps are windows into different depth levels of the submerged geosphere. In global geologic settings the sources of seep fluids range from 10s of meters (groundwater aquifers) to 10s of km (subducted oceanic plates) below the seafloor. At the sedimented active and passive margins organic-rich and evaporite-containing strata determine the final fluid composition, emitting characteristically gashydrate-derived methane and brine-associated non-methane hydrocarbons, respectively. Seeps transport dissolved and gaseous phase compounds to the ocean and sustain oasis-type ecosystems at the seafloor by providing bioactive reductants sulfide, methane and hydrogen. The interaction between fluid composition, fluxes and biota results in a diagnostic hydrocarbon-metazoan-microbe-carbonate association of which currently >80 are active globally. The single most important reaction is anoxic oxidation of methane by Archaea (AOM) with secondary reactions involving thiotrophy and carbonate mineral precipitation. Function, structure and composition of AOM-consortia and metazoan assemblages in concert with the characterization of biomarkers are overwhelming topics of seep research with increasingly frequent identification of ancient seep sites (>300 million years ago). The library of biomarkers as well as fossilized microbial bodies grows steadily aided by the fortuitous situation that both are preserved in carbonate precipitates. The Gulf of Mexico, the Black Sea and the eastern Mediterranean Sea are sites of classical and ongoing seep studies. Large-scale new studies of seeps by the national consortia have been initiated in the Indian Ocean, the South China Sea and the Eastern Sea off Korea where gas hydrate layers are being drilled in the search for new energy.

1

Introduction

The deep hydrosphere amounts to about 20 million cubic km of water, comprised of pore waters of marine sediments, hydrothermal fluids of the fractured underlying oceanic crust, and fluids of the upper mantle. This corresponds to about 4/5 of the water locked in all the polar ice on Earth and considerably exceeds the amount of groundwater on all its continents (Suess and Linke, 2006; Wallmann, 2001). Conceptually, the deep hydrosphere is analogous to the deep biosphere, where the total biomass also exceeds other biomass reservoirs on the Earth’s surface (Lipp et al., 2008). Important windows into different depth levels of this geosphere are provided by marine cold seeps. They are transport pathways for dissolved constituents to the ocean and sustain unique oasis-type ecosystems at the seafloor. Seeps are not necessarily at ambient seafloor temperature but more characteristically flow slowly in comparison to hydrothermal vents. Long-standing objectives of seep studies have been the quantification of water and methane transfer, characterization of source-depth, and the role of seep transport in global elemental budgets and elemental recycling (Wallmann, 2001; Plank, 2002). Global tectonic settings provide an enormous range of levels of depth for fluid generation. For example, seeps range from effluents of shallow groundwater aquifers, underlying the shelves by a few 10s of meters, to waters expelled through oceanic fore-arcs, generated 10s of km below from subducted tectonic plates. The return flow at active margins is generally even more deeply sourced than at passive margins. Characterization cannot be based on margin tectonics alone, however, because reactions with host-materials may overprint the seep fluid composition during upward flow, the tectonic environment is the driving force and a first order factor to be considered for understanding seeps. Fluid characteristics and seep environment resulting

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from the interplay with biota will be discussed next and important issues as they relate to seep microbiology will conclude the topic.

2

Seeps at Active Plate Margins

2.1

Oceanic Plate and Continental Plate Convergence

By far the most frequent and best studied seafloor seeps worldwide occur at the convergence between oceanic and continental plates and extend from the upper continental slopes to the deep trenches (> Fig. 1). Indeed, it was research at global deep-sea trench systems that first revealed the uniqueness of seeps and the mechanism of large-scale fluid expulsion was recognized as dewatering of sediments (Kulm et al., 1986; LePichon et al., 1987). The dewatering occurs in response to a lateral compression from plate movement. Sedimentladen, oceanic plates move under less dense continental plates. Sediments are either scraped off and accreted onto the edge of the overriding plate or are passed by at its base and become subducted. Off-scraping and by-passing lead to two types of subduction margins – accretionary and erosive – which constitute the global deep-sea trench system and generate seep fluids from different depths (LePichon et al., 1993; Moore and Silver, 2003; Shipley et al., 1990; von Huene et al., 2004). Accretionary margins consist of a series of ridges oriented parallel to the trench axis. In landward troughs between the ridges, known as fore-arc basins, thick sediments are deposited. The ridges as well as the basins are the source of seep fluids. At erosive margins small accretionary prisms can develop near the trench, but sediment and oceanic crust is carried downward with the plate, thereby removing material from or adding to the underside of the overriding plate. As a consequence, the margin subsides, fractures, and eventually the plate’s edges are destroyed (> Fig. 2). It is near the front where seeps initially form. Farther under the overriding plate, increasing temperatures and higher pressures release mineral-bound water by dehydration, forcing large amounts of fluids upward through fractures. Fluids mixed with mud form mud mounds and seeps, preferentially aligned above subsurface isotherms (Ranero and von Huene, 2000; Sahling et al., 2008). The destructive action at the plate’s edge is particularly severe when volcanic seamounts are subducted. When these elevated basaltic features arrive at the trench, riding on the oceanic crust, they plough into the continent leaving scars and scarps. Ensuing faulting and bulging of the sediments greatly facilitate fluid escape and seep formation.

2.2

Oceanic Plate and Oceanic Plate Convergence

Emission of deep-sourced fluids containing a biotic methane and hydrogen, results from convergence of two oceanic plates. The down-going plate contains little sediment, instead its top undergoes dehydration as temperature and pressure increase. This may take place as deep as 30 km below the seafloor. The liberated fluids then ascend and alter the overlying plate through hydration, forming serpentine from common olivine. The alteration products are carried upward, along with the excess of fluids, and exit as seeps from serpentinite mud mounds. This scenario was first recognized at the Mariana Trench (Fryer et al., 1995; Mottl et al., 2004).

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. Figure 1 Seep locations with hydrocarbon-metazoan-microbe-carbonate characteristics; active margin sites (blue), passive margin sites (orange) including groundwater seeps; sites at transform margins (green). Distribution based on first seep location maps by Campbell et al. (2002) and Campbell (2006) with complete site references. New sites are from Coleman and Ballard (2001), Jeong et al. (2004), Loncke et al. (2004), Olu et al. (2004), Judd et al. (2007), Mastarlerz et al. (2007), Hovland (2007), Han et al. (2008), Pape et al. (2008), Sahling et al. (2008), Geli et al. (2008), Hilario and Cunha (2008), Sellanes et al. (2008); dead seep clam site off India from Collett et al. (2008); revisited sites from Sahling et al. (2008), Olu-Le Roy et al. (2007a, b), Naudts et al. (2006), Pape et al. (2008). Sites based on dredged tube worms (triangles), mostly from deep-sea trenches, are from Ivanov (1963) prior to recognition that seep communities at plate boundaries are sites of hydrocarbon seepage. Up-dates of new site locations are found in Zhang and Lanoil (2004), Garcia-Gil and Judd (2007).

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. Figure 2 During plate convergence at erosive margins small accretionary prisms form at the trench, generating seeps similar to accretionary margins in response to squeezing of water-rich sediments. This is followed by dehydration releasing mineral-bound water, which is forced upward through hydro fractures, sometimes mixed with mud. Ejected products form mud mounds and seeps. Subducted volcanic seamounts contribute to the destruction of the overriding plate, leaving scars, scarps, faults, and bulges all facilitating fluid escape; modified from Ranero and von Huene (2000).

2.3

Transform Plate Boundaries

Transform boundaries generate prolific cold seeps where they crosscut thick sedimentary sequences providing fluid pathways for expulsion along the fault planes. At transform boundaries two plates slide past each other with only horizontal movement between them, posing enormous earthquake hazards depending on the configuration of the plate margins. Seeps at transform boundaries have not previously been considered in their own right, but because of the interest in earthquake precursors the focus has shifted to gas seepage at such tectonic settings (Ge´li et al., 2008).

3

Seeps at Passive Continental Margins and Shelves

On passive margins the variety of geologic settings, the different mechanisms of fluid expulsion, and the worldwide occurrence of cold seeps are immense (> Figs. 1 and 3). Pockmarks on shelves and slopes are expressions of seeps fed from submerged aquifers, from over-pressured formations containing hydrocarbons and brines, or from rapidly deposited accumulations of water-rich sediments, as in deltas. Hydrocarbon seeps have long guided offshore exploration for oil and gas deposits (Judd and Hovland, 2007).

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Greenhouse gas emission Tube worms Microbes Bivalves

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. Figure 3 Seep manifestations with hydrocarbon-metazoan-microbe-carbonates at sedimented margins, typically passive margins (adapted from Moore 1999; Suess and Linke 2006). Groundwater seeps are found in coastal areas and brine pools, liquid hydrocarbon seeps, above subsurface salt domes and at any water depths from 100s to >3000m. Methane reefs at mid-slopes result from AOM-generated carbonate mineral precipitation. Methane hydrate rafts-infrequently observedtransfer methane directly from the seafloor to the atmosphere.

Groundwater seepage from sub-seafloor extensions of aquifers has been known since the early days of seafarers (Sonrel, 1868). Today groundwater seepage carries pesticides, herbicides, and fertilizer residues into shelf waters of coastal areas. Pumping for drinking water depletes groundwater reservoirs and allows seawater to enter, or high tidal ranges force it back into the aquifers. The result is widespread salt invasion of groundwater in coastal areas and other problems regarding global groundwater availability and usage (Gallardo et al., 2006). The driving mechanisms for fluid expulsion at passive margins are loading by sediment, differential compaction, overpressure and facies changes. Hence, rapidly accumulating waterrich sediments generate seeps and mud mounds in deltas, deep-sea fans, and along passive margins facing the open ocean as well as in marginal seas. In short, any changes involving different permeabilities of fluid-rich strata, such as ash layers, turbidites, sands and silts, drift sediments, sand and silts; even buried reefs when intersected by faults open up pathways for fluid escape.

4

Common Seep Characteristics at Active and Passive Margins

The type of fluids expelled back into the ocean at accretionary, erosive and transform margins, or at passive margins -whether deeply- or shallow-sourced -depend largely on the following factors: thickness of the sediment packet, rate of sedimentation, age, cooling history, and the morphology of moving or stationary plates. Organic-rich and evaporite-containing strata are end-members in determining the final seep fluid composition.

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Sedimented Margins

Fluids expelled through seeps at sedimented margins contain the remineralized nutrients (silica, phosphate, ammonia and alkalinity) and hydrogen sulfide, as well as dissolved and free methane from microbial degradation of sedimentary organic matter. Thick sediments often are deposited along continental margins associated with coastal upwelling or otherwise high primary productivity in response to nutrient loading from nearby continents. Such sediments are rich in organic matter, accumulate rapidly and thus greatly favour sulphate reduction and methanogenesis of the microbially-mediated early diagenetic reaction sequence. Most sedimented margins generate enough biogenic methane to move upwards, exiting as dissolved or free gas directly into the bottom water or alternatively reaching the methane-hydrate zone where it is retained in layers of gas hydrate (> Chapter 3, Vol. 1, Part 1). While free gas ascends through seismically-anomalous conduits (acoustic blanket zones; gas chimneys) and forms acoustically detectable plumes in the water column, gas-hydrate layers are detected seismically as bottom simulating reflectors (BSRs). Currently these layers are being drilled worldwide in the frantic search for new and cleaner energy (https:circle.ubc.ca/handle/2429/1022).

4.1.1

Formation and Manifestations of the Hydrocarbon-MetazoanMicrobe-Carbonate Association

Processes leading to formation of the diagnostic association, as presently known, are illustrated in > Fig. 4 (modified from Suess and Linke, 2006). Bioirrigation and bubble-induced mixing at seeps (Haeckel et al., 2007) drive the interactions between methane, AOM-activity, macro-biota and formation of carbonates. Methane, either from subsurface gas hydrate, from ascending bubbles or in the dissolved gas phase, supplies the AOM-consortia which aggregate at different sub seafloor depths (Reaction 1). They generate hydrogen sulfide which rises and is oxidized either in microbial mats at the surface or within the macro fauna by symbionts

Oxygen Sulfate Nitrate Sulfate Oxygen

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. Figure 4 Different flux rates (arrows) supply methane to AOM-consortia (red–green circles) which consume seawater sulfate; rising hydrogen sulfide is oxidized by seafloor microbial mats or by macrofaunae symbionts, using oxygen or nitrate. In the process, calcium carbonate precipitates.

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using oxygen or nitrate (Reactions 2a and 2b). Bivalves pump oxygen downward whereas tubeworms (not shown in > Fig. 4) extract hydrogen sulfide through their roots (Freytag et al., 2001). As a consequence of the AOM-activity, calcium carbonate phases are precipitated (Reaction 3). Earlier views favoured accidental reaction from the by-product of AOM, now it is thought plausible that the microbial community may actively promote precipitation even of selected mineral phases (van Lith et al., 2003).

4.1.2

Gas Hydrates

Gas hydrate forms from the methane-supersaturated pore fluids at elevated pressure (>60 bar) and low temperature ( Chapter 3, Vol. 1, Part 1)). The top of the natural gas hydrate stability zone (GHSZ) along the global ocean margins starts at water depths as shallow as 300 m in frigid Arctic bottom waters and around 750 m in warm subtropical bottom waters. The base of the GHSZ varies with the water depth and the geothermal gradient and may range from 100s to 1,000s of meters below sea floor (Paull and Dillon, 2001; Suess, 2003). Methane usually originates from fermentative decomposition of organic matter or is produced by bacterial CO2 reduction. In some settings gas hydrate forms from thermogenic methane which migrate from deeper hydrocarbon reservoirs (MacDonald et al., 2004; Sassen et al., 2004). These sources can readily be distinguished from one another by stable C- and D/H-isotopes of CH4 (Whiticar, 1999). In seep waters high precision Cl-analyses, coupled with O- and D/H-isotopes of H2O indicate the presence of gas hydrates in the subsurface which may drive seep flow (Haeckel et al., 2004; Hesse and Harrison, 1991).

4.1.3

Anoxic Oxidation of Methane

Subsurface gas hydrates and ascending free methane at seeps provide the C-reservoir for the AOM-consortia consuming interstitial SO4 (Boetius et al., 2000; Orphan et al., 2001). The gradient of SO4 within the near-surface sediments is a reliable indicator for rapid upward flow of methane-rich seep fluids. Gradients as steep as 1 mmol cm 1 often occur directly beneath microbial mats (Knittel et al., 2003). The sub-surface depth of the sulphate-methaneinterface has become a standard indicator for mapping seep locations and exploring for gas hydrates (Borowski et al., 1996). Function, structure and composition of AOM-consortia in concert with biomarker characterization are currently the overwhelming topics of seep research. Biomarkers are highly depleted in 13C relative to their carbon source and are linked to methanotrophic archaea (Elvert et al., 1999; Hinrichs et al., 1999). This results in isotopic compositions as low as 137‰ PDB (i.e., hydroxyarchaeol; Birgel et al., 2008a) depending on whether biogenic or thermogenic methane is utilized. The inexhaustible reservoir of methane carbon available to the consortia in seep environments maximizes the kinetic carbon isotope effect (Elvert et al., 2001). Biomarkers from AOM-consortia are identified from sediments and authigenic carbonates of recent seep sites and increasingly from deposits of ancient seeps (Campbell et al., 2002; Campbell, 2006). So far, the oldest biomarker record was found in a 300 million year-old limestone (Birgel et al., 2008b). The library of biomarkers grows steadily

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with different classes of compounds, as for example biphytanic diacids (Birgel et al., 2008a) and extended hydroxyarchaeol (Stadnitskaia et al., 2008), as well as archaeal and bacterial intact polar lipids (Rossel et al., 2008).

4.1.4

Authigenic Carbonates

A direct consequence of AOM is the precipitation of carbonate minerals. The association of carbonates with gas emissions, pockmarks, and populations of chemosynthetic faunas is the most widely used criteria to identify seeps. Indeed, the carbonate build-ups at seep sites have become known as chemoherms (Aharon, 1994). The dominant mineral phases are aragonite, Mg-calcite, and (proto)-dolomite but also Fe- and Mn-carbonates occur. Their d13C and d18O composition ranges from 60 to 30 ‰ PDB and +8 to +3 ‰ PDB, respectively, depending on the C- and O-source, the temperature, and the specific mineral phases being formed (Aharon, 2000; Bohrmann et al., 1998; Chen et al., 2005; Ferrel and Aharon, 1994; Greinert et al., 2001; Han et al., 2004, 2008; Ritger et al., 1987; Reitner et al., 2005a, b; Teichert et al., 2005). Hydrate water, dehydration water, glacial-interglacial seawater and – in rare cases – meteoric water determine the O-isotope make-up; whereas biogenic, thermogenic, and abiotic methane carbon, becoming strongly fractionated during AOM, determine the eventual C-isotope signal. The stable isotopes and the mineralogies are robust criteria, along with biomarker, for recognizing ancient seeps.

4.1.5

Biota

Numerous reviews and case studies (Cordes et al., 2007; Levin 2005; Levin et al., 2007; Olu-Le Roy et al., 2007a, b; Sahling et al., 2002; Sibuet and Olu, 1998) summarize vent taxa, document chemoautothrophy, energy flux, relate community structure to seep intensity (Sahling et al., 2003), and model interactions between macro-fauna (tubeworms) and microbial consortia (Cordes et al., 2005; Luff et al., 2004). Interestingly, tube worms, initially thought of as indicator organisms for abyssal environments (Ivanov, 1963) remained unrecognized as members of the seep fauna until their distribution in deep-sea trenches could be related to convergent plate boundaries (Suess et al., 1985). New chemosynthetic species are continuously recognized at seeps (Reimer et al., 2007; Hilario and Cunha 2008; Sellanes et al., 2008).

4.1.6

The Black Sea

The Black Sea has been accorded a special place in the ongoing seep studies for its anoxic, methane-rich water column below about 100 m and its thick, up to 16 km, organic-rich sediments. Over 2,700 seeps were mapped off Bulgaria and the Ukraine (Naudts et al., 2006). The sites are concentrated at the shelf-slope break extending downslope to about 725 m. In the sub-seafloor below that water depth the stability limit of methane hydrates has been projected from the water temperature at the bottom and the geothermal gradient, implying that above 725 m gaseous methane escapes into the water at the bottom and the methane below is retained as hydrate. The seep province continues off the Crimean Peninsula, where in the eastern basin at 1,000–2,300 m, gas hydrates, methane plumes in the water column and mud mounds were discovered (Greinert et al., 2006) (> Chapter 13, Vol. 1, Part 3). Still

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farther east at the margin off Georgia, extensive methane seepage is found in the Batumi area (Bohrmann et al., 2008). Throughout the Black Sea authigenic carbonates of many morphologies and compositions are associated with the seep provinces although seep macro fauna is absent. Instead microbial communities, colonies, mats, and microbial reefs abound (Michaelis et al., 2002; Peckmann et al., 2001; Reitner et al., 2005a, b; Schubert et al., 2006; Thiel et al., 2001; Treude et al., 2007).

4.2

Evaporitic Host-Strata: The Gulf of Mexico and The Eastern Mediterranean Sea

Where underlain by evaporites, loading by sediments causes salt tectonics, e.g. the formation of salt domes and salt ridges. Here low density and low viscosity salts escape the pressure of overburden by flowing upward. Salt domes push through the overlying strata, dragging them upwards, and cause faults along their flanks. This facilitates migration of fluids. The tops of salt domes, near the seafloor, often dissolve away by circulating seawater and collapse. The result is large pockmarks, resembling the circumference of the underlying salt dome and usually containing brine pools and significant amounts of non-methane hydrocarbons including asphalt. The shelves and slopes surrounding the Gulf of Mexico contain such seeps and pockmarks related to salt tectonics as does the seafloor of the slope off Yucatan. These are important provinces of oil and gas exploration and of classical hydrocarbon seep studies (Brooks et al., 1987; Kennicut et al., 1985; MacDonald et al., 1990, 2004; Sassen et al., 2004). The Messinian salt underlying the Mediterranean Sea affects seeps at the easternmost end below the sediments of the Nile deep-sea fan (Loncke et al., 2006). Seeps, mud mounds, methane plumes, carbonates and microbes have been reported (Dupre´ et al., 2007; Gontharet et al., 2007). Farther west, beneath the Mediterranean Ridge, evaporites provide the source material for brine expulsion. At 2,000–3,000 m brine pools and mud mounds form the seafloor (Westbrook and Reston, 2002). The brine composition depends on the depths within the salt sequence from which the fluid is generated. It is dominated by sodium chloride, magnesium chloride and potassium chloride (Wallmann et al., 1997). While the mud mounds show the complete hydrocarbon-metazoan-microbe-carbonate association (Olu et al., 2004) nothing is known about microbial life in and around the deep-water brine pools of such as extraordinary composition.

5

Deep Fluids from Erosive and Transform Margins

There are no previous attempts known to characterize the seep fluids emitted at erosive and transform margins in their own right, although some specific properties stand out. As higher temperatures and pressures and portions of the down-going plate are involved, mineralbound water from opal or clays and elements susceptible to high pressure and temperature mobilization are added to the upward flow. Mineral-bound water dilutes the salinity of the seep fluid, is characteristically 18O-enriched and contains such mobilized elements as boron and lithium (Chan and Kastner, 2000; Da¨hlmann and deLange, 2003; Hensen et al., 2004). At still higher pressures and temperatures serpentinization sets in. During these reactions methane is formed from reduction of oceanic carbon dioxide by hydrogen which in turn is set free from common olivine reacting with seawater (Sleep et al., 2004, > Chapter 14, Vol. 1, Part 3). Free hydrogen that is not consumed by CO2-reduction is abundant at

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serpentinization-driven seeps as are Archaea thriving at pH-values of >12 (Mottl et al., 2003). Mud mounds are dominant seafloor manifestations of deep-sourced seeps at erosive margins. At the Mariana Fore-Arc and the Costa Rica margin they are aligned above sub-seafloor isotherms (Mottl et al., 2004; Ranero and von Huene, 2000). Hence their geochemical and isotopic characteristics provide good evidence for depths of fluid generation. At mud mounds not just fluids are expelled but also material from the conduit walls. These products may transport microbial life from the deep biosphere to the seafloor. Fluids from transform margins are derived from a great depth because steeply dipping and deep-reaching fault planes are common for the transform fault systems. The Ba-enrichment at seeps observed at the San Clemente transform (McQuay et al., 2008) and the Sakhalin (Greinert et al., 2002) transform faults is noteworthy although not fully understood. Perhaps microbial SO4-reduction at depth, where pore water sulphate has been exhausted, utilizes particulate sedimentary barite (BaSO4). This would generate high concentrations of barium which in turn are expelled at the cold seeps where it would reprecipitate with seawater sulfate. This potentially important process involving the deep biosphere awaits further research.

6

Research Needs: Significant Issues of Seep Research

6.1

Global Methane Emissions from Seeps

Global assessment of seep fluxes must consider the role of the benthic filter (seafloor biota) in modulating methane output and hence reducing fluxes to the ocean (Boetius and Suess, 2004; Jørgensen and Boetius, 2007; Niemann et al., 2005; Sommer et al., 2006). Seep fluxes may contribute up to 15% of the natural methane sources in the atmosphere (Judd et al., 2002). The lack of appropriately identified geosphere methane in the current climate models is apparent. To improve this situation, fluid flow and bubble escape need to be quantified and up-scaled. Deep-sea landers equipped with seep meters deployed at single sites (Tryon et al., 1999) recorded flow rates between 102 cm y 1. A new acoustic technique, GasQuant, based on the backscatter intensity of bubbles, scans escaping plumes horizontally at the sea floor and integrates bubble-spectra for total gas fluxes (Greinert, 2008). Time- and space-integrated estimates, a geophysical approach in translating seismic velocity changes from loss of pore space of subducted sediments into fluid emission (Saito and Goldberg, 2001; von Huene et al., 1998), should all be pursued further and refined.

6.2

Ancient Seeps and Fossilization of Microbial Structures

Documentation of microbial activity from ancient seeps progresses well. The geologic time scale for seep activity is steadily pushed back with ancient plate boundaries and passive margins being recognized as characteristic settings (Campbell et al., 2002; Campbell, 2006). Sediment fabric and organic geochemistry of geological material will continue to yield sites of ancient seeps. The concern over contamination of the geological sample materials is reduced since microbial bodies are fully encased in the carbonate matrix formed during AOM. Likewise, preparatory work in isolating biomarkers from authigenic carbonates ascertains contamination-free samples. New miniaturized bio-signature extraction procedures have recently been introduced (Leefmann et al., 2007). Abundant structures of suspected fossilized microbes are present in seep carbonates (Peckmann et al., 2004). Among these are

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filamentous, rod-shaped, cylindrical, spindle- and dumb-bell shaped bodies that await identification (Han et al., 2008). Fossilized bodies, resembling microbial morphologies and consisting solely of AOM-carbonates, e.g. aragonite, high Mg-calcite, and dolomite grains, and their biomarker make-up will provide considerable new knowledge.

6.3

Benthic Microbial Fuel Cell at seeps

New benthic microbial fuel cell technology (BMFC) might some day power instruments needed to arrive at global methane emissions from the seafloor. Electrical power produced at seeps to run deep-sea monitoring instruments in place of batteries, is no longer a distant vision (Nielsen et al., 2007, 2008). Seep-sourced sulfide, methane, hydrogen and dissolved organic matter are bioreactive reductants that represent a natural reservoir for chemical energy from the geosphere. If seep sites are stable and flow rates are maintained, they should supply these species to the sediment water-interface continuously, maintaining the steep redox gradient needed to produce electrical power in situ (Reimers et al., 2006). Benthic microbial fuel cells employ inert electrodes of large surface areas to couple the microbial oxidation of reduced species derived from sediments to the reduction of oxygen dissolved in the bottom water. Advanced fuel cell design and analysis of microbial bio-films from both anodes and cathodes should optimize power production (Nielsen et al., 2007) and clarify the molecular mechanisms of extra cellular electron transfer used by microbes. Electrical power >140 mW m 2 has been sustained during a prototype BMFC deployment for over 120 days at a transform fault seep in Monterey Canyon off California (Nielsen et al., 2007). Up-scaling with power conversion to higher voltages is possible.

Acknowledgments Special thanks goes to colleagues who provided information, material and general advice during preparation of this chapter. Kathleen A. Campbell (Geology and Environmental Sciences, University of Auckland) provided me with an extensive compilation of seep sites; Jens Greinert (Netherlands Institute for Sea Research, Den Burg) helped with making available the global tectonic map; Ce´sar R. Ranero (Instituto de Ciencias del Mar, Barcelona) generated the basic cartoon on erosive margin dewatering (> Fig. 2); Marcus Elvert and Daniel Birgel (Geowissenschaften, Universita¨t Bremen) updated the biomarker information, and Clare E. Reimers (College of Oceanic and Atmospheric Sciences, Oregon State University) drew my attention to the Benthic Microbial Fuel Cell technology. Zona Suess helped, as always, with the intricacies of the English language. This is contribution GEOTECH-868.

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of Abyssoanthus nankaiensis, a new family, new genus and new species of deep-sea zoanthid (Anthozoa:Hexacorallia: Zoantharia) from a north-west Pacific methane cold seep. Invertebr Syst 21: 255–262. Reimers CE, Girguis P, Stecher H III, Ryckelynck N, Tender LM, Whaling P (2006) Microbial fuel cell energy from an ocean cold seep. Geobiology 4: 123–136. Reitner J, Peckmann J, Blumenberg M, Michaelis W, Reimer A, Thiel V (2005a) Concretionary methaneseep carbonates and associated microbial communities in Black Sea sediments. Palaeogeogr Palaeoclimatol Palaeoecol 227: 18–30. Reitner J, Peckmann J, Reimer A, Schumann G, Thiel V (2005b) Methane-derived carbonate build-ups and associated microbial communities at cold seeps on the lower Crimean shelf (Black Sea). Facies 51: 66–79. Ritger S, Carson B, Suess E (1987) Methane-derivedauthigenic carbonates formed by subduction-induced pore-water expulsion along the Oregon/Washington margin. Geol Soc Am Bull 98: 147–156. Rossel PE, Lipp JS, Fredricks HF, Arnds J, Boetius A, Elvert M, Henrichs KU (2008) Intact polar lipids of anaerobic methanotrophic archaea and associated bacteria. Organic Geochem 39: 992–999. Sahling H, Masson DG, Ranero CR, Hu¨hnerbach V, Weinrebe W, Klaucke I, Bu¨rk D, Bru¨ckmann W, Suess E (2008) Fluid seepage at the continental margin offshore Costa Rica and southern Nicaragua. Geochem Geophys Geosyst 9:Q05S05. doi: 10.1029/ 2008GC001978. Sahling H, Galkin SV, Salyuk A, Greinert J, Foerstel H, Piepenburg D, Suess E (2003) Depth-related structure and ecological significance of cold-seep communities. A case study from the Sea of Okhotsk. Deep Sea Res 50: 1391–1409. Sahling H, Rickert D, Lee RW, Linke P, Suess E (2002) Macrofaunal community structure and sulfide flux at gas hydrate deposits from the Cascadia convergent margin; NE Pacific. Mar Ecol Progr Ser 231: 121–138. Saito S, Goldberg D (2001) Compaction and dewatering processes of the oceanic sediments in the Costa Rica and Barbados subduction zones: estimates form in situ physical property mneasurements. Earth Planet Sci Lett 191: 283–293. Sassen R, Roberts HH, Carney R, Milkov A, DeFreitas DA, Lanoil B, Zhang CL (2004) Free hydrocarbon gas, gas hydrate and authigenic minerals in chemosynthetic communities of the northern Gulf of Mexico continental slope: relation to microbial process. Chem Geol 205: 195–217. Schubert CJ et al. (2006) Methanotrophic microbial communities associated with bubble plumes above

gas seeps in the Black Sea. Geochem Geophys Geosyst 7: Q04002. doi: 10.1029/2005GC001049. Sellanes J, Quiroga E, Neira C (2008) Megafauna community structure and trophic relationships at the recently discovered Concepcio´n methane seep area, Chile, 36degS. ICES J Mar Sci Adv Access 65: 1102–1111. doi: 10.1093/icesjms/fsn099. Shipley TH, Stoffa PL, Dean DF (1990) Underthrust sediments, fluid migration paths, and mud volcanoes associated with the accretionary wedge off Costa Rica: Middle America Trench. J Geophys Res 95(B6): 8743–8752. Sibuet M, Olu K (1998) Biogeography, biodiversity and fluid dependence of deep-sea cold-seep communities at active and passive margins. Deep Sea Res Part II 45: 517–567. Sleep NH, Maibom A, Fridriksson Th, Coleman RG, Bird DK (2004) H2-rich fluids from serpentinization: geochemical and biotic implications. Proc Natl Acad Sci 101: 12818–12823. Sommer S, Pfannkuche O, Linke P, Luff R, Greinert J, Drews M, Gubsch S, Pieper M, Poser M, Viergutz T (2006) Efficiency of the benthic filter: biological control of the emission of dissolved methane from sediments containing shallow gas hydrates at Hydrate Ridge. Global Biogeochem Cycles 20: GB2019. Sonrel L (1868) Le Fond De La Mer. In: Bibliotheque des Merveilles, Charton MA (ed.). Librairie de la Hachette et Cie., Paris, Blvd. Saint-Germain No. 77, 540 pp. Stradnitskaia A, Bouloubassi I, Elvert M, Hinrichs KU, Sinninghe Damste´ JS (2008) Extended hydroxylarchaeol, a novel lipid biomarker for anaerobic methanotrophy in cold seep habitats. Org Geochem 39: 1007–1014. Suess E (2002) The evolution of an idea: from avoiding gas hydrates to actively drilling for them JOIDES J 28: 45–50. Suess E, Carson B, Ritger SD, Moore JC, Jones ML, Kulm LD, Cochrane GR (1985) Biological communities at vent sites along the subduction zone off Oregon. Biol Soc Washington Bull 6: 475–484. Suess E, Linke P (2006) Der Ozean unter dem Meeresboden – Kalte Quellen als Oasen der Tiefsee. In Expedition Erde, 2nd edn. Wefer G (ed.). New York: Springer, pp 88–101. Teichert BMA, Bohrmann G, Suess E (2005) Chemoherms on Hydrate Ridge – unique microbiallymediated carbonate build-ups growing into the water column. Palaeogeogr Palaeoclimatol Palaeoecol 227: 67–85. Thiel V, Peckmann J, Richnow HH, Luth U, Reitner J, Michaelis W (2001) Molecular signals for anaerobic methane oxidation in Black Sea seep carbonates and a microbial mat. Mar Chem 73: 97–112.

Marine Cold Seeps Treude T et al. (2007) Consumption of methane and CO2 by methanotrophic microbial mats from gas seeps of the anoxic Black Sea. Appl Environ Microbiol 73: 2271–2283. Tryon MD, Brown KM, Torres ME, Trehu AM, McManus J, Collier RW (1999) Measurements of transient and downward fluid flow near episodic methane gas vents, Hydrate Ridge, Cascadia. Geology 27: 1075–1078. van Lieth Y, Warthmann R, Vaconcelos C, McKenzie JA (2003) Microbial fossilization in carbonate sediments: a results of bacterial surface involvement in dolomite formation. Sedimentology 50: 237–245. von Huene R, Klaeschen D, Gutcher M, Fru¨hn J (1998) Mass and fluid flux during accretion at the Alaska margin. Geol. Soc. Am., Bull. 110: 468–482. von Huene R, Ranero CR, Vannucchi P (2004) Generic model of subduction erosion. Geology 32: 913–916.

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Wallmann K (2001) The geological water cycle and the evolution of marine d180 values. Geochim. Cosmochim Acta 65: 2469–2485. Wallmann K, Suess E, Westbrook GH, Winckler G, Cita MG (1997) Salty brines on the Mediterranean sea floor. Nature 387: 31–32. Westbrook GH, Reston TJ (2002) The accretionary complex of the Mediterranean Ridge: tectonics, fluid flow and the formation of brine lakes – an introduction to the special issue of Marine Geology. Mar Geol 186: 1–8. Whiticar MJ (1999) Carbon and hydrogen isotope systematics of bacterial formation and oxidation of methane. Chem Geol 161: 291–314. Zhang CL, Lanoil B (guest eds.) (2004) Geomicrobiology and biogeochemistry of gas hydrates and hydrocarbon seeps. Chem Geol 205: 187–194.

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13 Mud Volcanoes H. Niemann1,2 . A. Boetius1,3 1 Max Planck Institute for Marine Microbiology, Bremen, Germany 2 Institute for Environmental Geosciences, University of Basel, Basel, Switzerland 3 Alfred Wegener Institute for Marine and Polar Research, Bremerhaven, Germany [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 206 2 Hydrocarbon Emissions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 208 3 Geochemical Forcing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 209 4 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 211

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_13, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Mud volcanoes are frequently encountered geo-structures at active and passive continental margins. In contrast to magmatic volcanoes, mud volcanoes are marine or terrestrial, topographic elevations built from vertically rising fluidized mud or mud breccia. Commonly, these structures have a crater, hummocky rime and caldera. Mud volcanism is triggered by various geological processes which lead to a high pore fluid pressure at great depth, sediment instabilities and a subsequent discharge of mud, fluids and gases such as hydrocarbons (mostly the greenhouse gas methane). Although global estimates of methane emissions from mud volcanoes vary over two orders of magnitude, mud volcanism could be an important source for atmospheric methane. However, a substantial fraction of the hydrocarbons are retained in the mud volcanoes surface sediments. Here, the upwelled hydrocarbons fuel a variety of free-living and symbiotic, chemosynthetic communities that oxidize these with electron acceptors such as oxygen or sulphate from the water column or the atmosphere. The activity of the chemosynthetic communities is regulated by the availability of either electron donors (hydrocarbons) or acceptors which, in return, is determined by mass transport processes. Most important in this context are the magnitudes of upward advection of electron donors and the influx of electron acceptors due to diffusion and bioirrigation.

1

Introduction

Mud volcanoes are geological structures bearing only little morphological resemblances to magmatic volcanoes. In contrast to true volcanoes which expel magmatic material at plate boundaries and mantle plumes (Schmincke, 2006), mud volcanoes are formed by vigorous mud discharge that is often accompanied by fluid and gas emissions commonly originating from a deep subsurface sedimentary sequence (Brown, 1990; Kopf, 2002; Milkov, 2000). Mud volcanoes have a long tradition of scientific investigation and references were already made in historical documents (e.g., ‘‘Naturalis Historia’’ by Pliny the Elder, first century AD). Nevertheless, the diversity of mud volcano shapes as well geological causes responsible for their formation lead to a variety of definitions and synonymous terms such as mud volcano, mud pie, mud mound and gryphon (among others). Hereafter, a mud volcano is defined as a marine or terrestrial, topographic elevation built from vertically rising fluidised mud or mud breccia (a mud matrix with clasts). Mud volcanism is caused by various geological processes such as tectonic accretion and faulting, rapid burial of sediments due to slope failures (olistostromes) or high sedimentation rates, fluid emissions from mineral dehydration as well as (true) volcanic and earth quake activities (Brown, 1990; Dimitrov, 2002; Kopf et al., 2001, 2002; Mellors et al., 2007; Milkov, 2000). These processes can lead to an abnormally high pore fluid pressure and sediment instabilities and consequently lead to the extrusion of mud, fluids, and gases such as hydrocarbons and carbon dioxide to the sea floor or earth surface (usually through a central conduit; > Fig. 1). A crater or active centre, hummocky rim and surrounding caldera are common features of mud volcanoes. However, the shape of the edifice can range from amorphous mud pies to conical formations and their size varies from a few meters to kilometres in circumference and a few decimetres to hundreds of meters in height. Viscosity and density of the extruded material as well as the duration of eruption events and the development stage of the edifice were identified as major factors determining the shape of mud volcanoes (Lance et al., 1998; Murton and Biggs, 2003; Stewart and Davies, 2006). In general, flat structures are composed of comparably liquid mud matrixes, while high and cone

Mud Volcanoes

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Continental slope Methane plume Chemosynthetic comunities Humocky rim

a

b

Caldera Crater Central conduit

c

d

e

. Figure 1 Potential genesis of a gas emitting (marine) mud volcano. (a) High pore fluid pressure leads to the formation of mud breccia at great depth where also gases (mainly methane) are produced. (b) Overpressurised mud breccia and gases migrate along sediment instabilities to the seafloor (c) which is eventually breached (d) and upheaved. (e) Scheme of the Haakon Mosby Mud Volcano. At a ‘‘typical’’ mud volcano, mud and gases are transported through a central conduit and extruded in a crater region. The crater is surrounded by a hummocky rim of displaced sediment material. After an initial outburst and deflation of source material, a caldera (collapse-structure) surrounds the mud volcano. Surface sediments of mud volcanoes can support a wide range of free-living and symbiotic, chemosynthetic organisms which oxidize the upwelled hydrocarbons and hydrogen sulphide with oxidants such as oxygen, nitrate or sulphate from the water column. Giant sulphide oxidizing bacteria forming white mats on the sea floor and symbiotic tube-worms colonizing the sea floor in meadow- or bush-like aggregations are prominent examples of chemosynthetic communities, which are visible for the naked eye. a-d Source: ARCHIMEDIX, e Sabine Lu¨deling, MedienIngenieure Bremen.

shaped edifices are build of successively, superimposed flows of more viscous material. Mud volcanoes may thus erupt in regular or irregular time intervals or emit mud, fluids and gases continuously. In addition, they may also become inactive when the source of gas expansion and fluid flow stops (Planke et al., 2003) but also new structures evolve such as the terrestrial LUSI mud volcano in 2006 (Mazzini et al., 2007). Three types of mud volcano activity are distinguished (Dimitrov, 2003 and references therein): (1) Lokbatan-type: This type of mud volcanisms was named after the Lokbatan mud volcano, Azerbaijan. Lokbatan type mud volcanoes are characterized by violent outbreaks and long phases of dormancy. (2) Chikishlyar-type: Calm, relatively weak and continuous venting of gas, water and mud are typical for this type of mud volcano.

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(3) Shugin-type: This type of mud volcanism is transitional between the other types, characterized by long periods of weak activity interrupted by eruptive events. Dimitrov (2003) suggested that this type of mud volcanism is the most common. This distinction is based on terrestrial mud volcanism, which has been investigated for a comparably long time. In some cases, also historical documents can be used to infer the mode of activity (Aliyev et al., 2002). In contrast, most oceanic mud volcanoes were discovered and investigated in the last decade, when appropriate high-resolution geophysical tools became available to science which can resolve a few m of difference in height above or below ground. However, from the bathymetry it cannot be resolved what activity-type a particular mud volcano may represent. Eruptive events could be separated by extremely long periods of dormancy. In addition to the temporal heterogeneity of activity, visual investigation of mud volcanoes by towed video cameras, submersibles or remotely operating vehicles showed that marine mud volcanism is also spatially diverse (Niemann et al., 2006b; Sauter et al., 2006). In general, a mud volcano has an active centre above a central conduit which is usually marked by steep temperature gradients, and seepage rates decrease towards the periphery. However, the active centre may not always be the geographical centre and the activity may not follow a concentric arrangement. Our knowledge about mud volcanoes in general and specific structures in particular is therefore very sketchy.

2

Hydrocarbon Emissions

The processes leading to mud volcanism on the continents as well as at active and passive continental margins are generally related to fluid and gas flow. Subsurface muds and shales in mud volcano-hosting regions often contain high amounts of methane and other hydrocarbons of thermogenic and/or microbial origin. Consequently, mud flows can be accompanied by vigorous gas expulsions, which may even ignite in contact with the atmosphere in terrestrial systems (Charlou et al., 2003; Kopf, 2002; Milkov, 2000; Somoza et al., 2003). Good examples for violent gas emissions from such structures are the terrestrial Lokbatan and the deep water Haakon Mosby Mud Volcano. During the last outbreak in 2001, a flame of about 400 m height lasted for more than a day above Lokbatan mud volcano (Aliyev et al., 2002; Mukhtarov et al., 2003). At Haakon Mosby, a gigantic methane plume of about 600 m was visible on echosounder systems during several cruises and jets of methane emitted from the sea floor were observed during submersible dives (Sauter et al., 2006; Vogt et al., 1997). The annual methane discharge from Haakon Mosby was estimated with 8 – 35 Mmol (0.1 – 0.5 Gg) of which free gas accounted for 60–90% (Niemann et al., 2006b; Sauter et al., 2006). About 650 to 900 terrestrial mud volcanoes are known (Kopf, 2003) but global estimates for marine mud volcanoes range between 800 and 100000 (Dimitrov, 2002, 2003; Kopf, 2003; Milkov, 2000; Milkov et al., 2003). For submarine mud volcanoes, it is often not known if and when these structures emit methane. As a result, global assessments of methane emissions from mud volcanoes vary considerably. Recent estimates suggest that terrestrial and shallow water mud volcanoes contribute between 2.2 and 6 Tg yr
1 of methane to the atmosphere (Dimitrov, 2003; Milkov et al., 2003) and that 27 Tg yr
1 of methane may escape from deep water mud volcanoes (Milkov et al., 2003). Revised estimates of the total methane emission from mud volcanoes range between 35–45 Tg yr
1 (Etiope and Milkov, 2004), 30–70 Tg yr
1 (Etiope and Klusman, 2002), and – when using only known structures

Mud Volcanoes

13

and correcting for the size of the edifice – between 0.3 Tg yr
1 (Kopf, 2003) and 1.4 Tg yr
1 (Kopf, 2002). In comparison to the annual methane emissions to the atmosphere (535 Tg yr
1, Judd et al., 2002), mud volcanism may consequently be a significant source for atmospheric methane.

3

Geochemical Forcing

In surface sediments of mud volcanoes potential electron donors such as hydrocarbons and, after their biological conversion with sulphate, hydrogen sulphide from deeper sediment layers meet electron acceptors such as oxygen, nitrate/nitrite, oxidized metals and sulphate from the water column or the atmosphere. In such redox transition zones, mud volcanoes were found to support a wide range of free-living and symbiotic, chemosynthetic organisms utilizing the subsurface energy sources (also known as ‘‘geofuels’’; > Fig. 1). Thereby, chemosynthetic organisms reduce the efflux of reduced molecules to the hydro- and atmosphere (See > Chapters on hydrocarbon and sulphur oxidising microbes in Vol. 2 ‘‘MICROBIAL UTILIZATION OF HYDROCARBONS, OILS AND LIPIDS’’ of this edition) (Alain et al., 2006; Jørgensen and Boetius, 2007; Joye et al., 2005; Niemann et al., 2006a, 2006b; Olu et al., 1997; Omoregie et al., (in review)). The most important metabolic pathways are methanotrophy (anaerobic oxidation of methane – AOM, and aerobic oxidation of methane - MOx), anaerobic and aerobic degradation of hydrocarbons, thiotrophy (sulphide oxidation with oxygen or nitrate - SOx) and in some recently discovered systems also iron oxidation. The distribution of chemosynthetic communities strongly depends on the availability of electron donors and acceptors which in return is regulated by physical mass transport processes and biological activities (de Beer et al., 2006; Lo¨sekann et al., 2007; Niemann et al., 2006b). Advection accounts for the majority of upward transport of electron donors from deeper sediment layers, while diffusion and bioirrigation are responsible for most of the influx of electron acceptors from the atmosphere or the water column into the mud volcano sediments (> Table 1). Advective transport at mud volcanoes is in the form of mud, fluid and free gas flow (see section 1). Direct measurements of advection are scarce (Brown et al., 2005; Linke et al., 1994; Mazzini et al., 2007; Sauter et al., 2006). In particular, rates of free gas and mud flow are poorly resolved. Also, the effect of mud and free gas flow on the distribution of chemosynthetic communities is mostly unknown. Fluid flow rates, on the other hand, can be modelled from geochemical porewater gradients and heat flow measurements, which allows for a comparably high temporal and spatial resolution. Recorded values for fluid flow at active mud volcanoes are typically a few centimetres to several metres per year (> Table 1). Except for the spatial and temporal heterogeneity of mud volcano activity, advective pore water transport is a linear process and the advective flux (Ja), i.e., the amount of a pore water solute crossing a given area per time, is determined by the flow velocity (va) and the concentration (C) of the solute: Ja ¼ va C

ð1Þ

(Note that C has to be corrected for porosity -f) The underlying mechanism of diffusion is Brownian motion (Einstein, 1905), which, for biogeochemical reactions, can be simplified to the heat induced, non-directional movement of atoms/molecules in water. Diffusive transport can be illustrated by assuming two spatially separated entities in sediments or the water column with high and low concentrations of a

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. Table 1 Fluid flow velocities (va; in cm yr
1) at selected submarine mud volcanoes Structure

Location

va

Ref.

Haakon Mosby

Barents Sea

40–600*

1,2

Dvurechenski

Black Sea

8–25

3

Capt. Arutyunov

Gulf of Cadiz

10–15

4

Mound 12

East Pacific

10

5

Atalante

West Atlantic

1cm s
1 were observed

given solute. The dissolved atoms/molecules will move randomly between both units. But more atoms/molecules will move from the unit of high concentration than from the unit of low concentration. This consequently leads to a net transport to the unit of low concentration until both units are equal in concentration. From this simple example it is apparent that the concentration difference is an important factor determining diffusive flux. The second important factor is the net velocity of the movement. However, in contrast to the linear mode of advective flow, diffusion is random. A diffusing atom/molecule will not move in one direction but, in a simplified manner, forward and backwards. As a result and when considering a large number of atoms/molecules, the mean traveled net distance (L) increases only by the square root of time (t): pffiffiffiffiffiffiffiffiffi L ¼ 2Dt; ð2Þ where D, the diffusion coefficient, is a compound specific constant usually expressed in cm2 yr
1 (note that D has to be corrected for temperature (T) and f, e.g., Boudreau, 1997). Equation (2) has the very counterintuitive implication that the net velocity of diffusion (vd) decreases with increasing diffusion distance: vd ¼ L=t ¼ 2D=L

ð3Þ

Important electron donors and acceptors at mud volcanoes only need about a ms to travel a distance of 1 mm but already a day for 1 cm and some month for 10 cm (> Table 2)! The diffusive flux (Jd) is hence determined by the concentration difference (dC), the diffusion distance (dx) and D. Assuming steady state conditions, i.e., none of the factors determining the flux changes over the time period of measurement, Jd can be calculated according to Fick’s first law of diffusion (Berner, 1980; Boudreau, 1997; Fick, 1855): Jd ¼ D  dC=dx

ð4Þ

For short distances and high concentration differences (i.e., a steep concentration gradient dC= ), diffusion is an efficient transport mechanism. Diffusive transport of electron acceptors dx may thus balance a high advective flux of electron donors from below. However, the redox transition zone in this scenario will be close to the sediment surface where the concentration gradients of the electron acceptors are steep.

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Mud Volcanoes

. Table 2 Diffusion distance (L) in relation to diffusion time (t) and velocity (vd) of methane, sulbhate and oxygen in sediments at a ‘‘typical’’ submarine mud volcano (T3 C; f80%) Methane

Sulphate

Oxygen

L

t

vd

t

vd

t

vd

1 mm

0.8 ms

40 km yr
1

1.3 ms

24 km yr
1

0.7 ms

48 km yr
1

1 mm

13.2 min

40 m yr
1

22 min

24 m yr
1

11 min

48 m yr
1

18 h

4.8 m yr
1

76 d

0.48 m yr
1

21 yr

4.8 cm yr
1

2.1 kyr

4.8 mm yr
1

209 kyr

0.48 mm yr
1

209 Myr

48 mm yr
1

1 cm

22 h


1

4 m yr

10 cm

92 d

0.4 m yr

1m

25 yr

4 cm yr
1

10 m

2.5 kyr

36 h


1

4 mm yr


1
1

100 m

250 kyr

0.4 mm yr

1 km

25 Myr

40 mm yr
1

2.4 m yr


1
1

150 d

0.24 m yr

41 yr

2.4 cm yr
1

4.1 kyr

2.4 mm yr


1

413 kyr

0.24 mm yr

41 Myr

24 mm yr
1


1

The transport of electron acceptors due to bioirrigation activities is a known but poorly quantified phenomenon at marine mud volcanoes (Haese et al., 2006; Niemann et al., 2006b) and other types of cold seeps (Cordes et al., 2005; Haese, 2002; Treude et al., 2003). Many mud volcanoes host large populations of chemosynthetic megafauna such as tube worms and bivalves mining for sulphide and methane. Thereby, oxygenated and sulphate rich sea water is flushed through burrows into deeper sediment layers where it becomes available for free living chemosynthetic microbes. Furthermore, some thiotrophic tube worms are known to secret sulphate actively through posterior body parts to fuel sulphate reduction in the sediment. The flux via bioirrigation (Jb) is solely dependent on the faunal (pumping) activity as well as their extension into the sediment. Jb can be calculated from the concentration differences of non-reactive tracers such as e.g., silica or bromide (Haese, 2002; Haese et al., 2006; Wallmann et al., 1997): Jb ¼ ahðC0
Cx Þ

ð5Þ


1

where a is the non-local exchange coefficient (in yr , dependent of faunal community composition and density) which has to be modeled from pore water concentration profiles. h is the thickness of the zone in which the transport occurs and C0 and Cx are the concentrations of the tracer in the bottom water and at depth, respectively. The few estimates available to date indicate that fluid flow due to bioirrigation may be 2 – 3 orders of magnitude higher than the physical transport (Haese et al., 2006; Wallmann et al., 1997). Because of the different modes and magnitudes of transport, the redox transition zones are found at various depth in mud volcano sediments ranging from the sediment surface to meters below sediment surface. For sediments devoid of burrowing megafauna, the depth is determined by the velocity of upward fluid flow (de Beer et al., 2006).

4

Research Needs

Due to the high spatial and temporal variability of fluid flow at mud volcanoes, and the many questions remaining to the functioning and interaction of geophysical forces as drivers of mud

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volcanism, there are still many open question as to the trigger, sources and change of their activity and longevity. For submarine mud volcanoes an important issue is the relation between gas and fluid flow, heat transport and the formation/dissociation of gas hydrates as well as its consequences for the distribution and activity of faunal communities. One of the best studied mud volcanoes in this regard is the Haakon Mosby Mud Volcano, which has been chosen as a site for long term observation of geophysical and biogeochemical processes of mud volcanism. Specifically for terrestrial mud volcanoes, very little is known about the occurrence, phylogeny, ecology, and activity of chemosynthetic communities.

References Alain K, Holler T, Musat F, Elvert M, Treude T, Kruger M (2006) Microbiological investigation of methaneand hydrocarbon-discharging mud volcanoes in the Carpathian Mountains, Romania. Environ Microbiol 8(4): 574–590. Aliyev AA, Guliyev IS, Belov IS (2002) Catalogue of recorded eruption of mud volcanoes of Azerbaijan (for period of years 1810–2001). Baker: Nafta Press. Aloisi G, Drews M, Wallmann K, Bohrmann G (2004) Fluid expulsion from the Dvurechenskii mud volcano (Black Sea) - Part I. Fluid sources and relevance to Li, B, Sr, I and dissolved inorganic nitrogen cycles. Earth Planet Sci Lett 225: 347–363. Berner RA (1980) Early Diagenesis - A Theoretical Approach. Princeton, NJ: Princeton University Press. Boudreau BP (1997) Diagenetic models and their implementation: Modelling transport and reactions in aquatic sediments. Springer. Brown KM (1990) The nature and hydrogeologic significance of mud diapirs and diatremes for accretionary systems. J Geophysical Research-Solid Earth and Planets 95(B6): 8969–8982. Brown KM, Tryon MD, DeShon HR, Dorman LM, Schwartz SY (2005) Correlated transient fluid pulsing and seismic tremor in the Costa Rica subduction zone. Earth Planet Sci Lett 238(1–2): 189–203. Charlou JL, Donval JP, Zitter T, Roy N, Jean-Baptiste P, Foucher JP, Woodside J (2003) Evidence of methane venting and geochemistry of brines on mud volcanoes of the eastern Mediterranean Sea. Deep-Sea Res Part I-Oceanogr Res Pap 50(8): 941–958. Cordes EE, Arthur MA, Shea K, Arvidson RS, Fisher CR (2005) Modeling the mutualistic interactions between tubeworms and microbial consortia, Plos Biology 3(3): 497–506. de Beer D, Sauter E, Niemann H, Kaul N, Foucher JP, Witte U, Schluter M, Boetius A (2006) In situ fluxes and zonation of microbial activity in surface sediments of the Hakon Mosby Mud Volcano. Limnol Oceanogr 51(3): 1315–1331.

Dimitrov LI (2002) Mud volcanoes - the most important pathway for degassing deeply buried sediments. Earth-Sci Rev 59(1–4): 49–76. Dimitrov LI (2003) Mud volcanoes - a significant source of atmospheric methane. Geo-Marine Let 23(3–4): 155–161. Einstein A (1905) The motion of elements suspended in static liquids as claimed in the molecular kinetic theory of heat. Annalen Der Physik 17(8): 549–560. Etiope G, Klusman RW (2002) Geologic emissions of methane to the atmosphere. Chemosphere 49(8): 777–789. Etiope G, Milkov AV (2004) A new estimate of global methane flux from onshore and shallow submarine mud volcanoes to the atmosphere. Environ Geol 46(8): 997–1002. ¨ ber Diffusion. Poggendorf ’s Annalen der Fick A (1855) U Physik und Chemie 94: 59–86. Haese RR (2002) Macrobenthic activity and its effects on biogeochemical reactions and fluxes. In Ocean Margin Systems. G Wefer, D Billet, D Hebbeln, BB Jørgensen, M Schlu¨ter, and TCE Van Weering (eds.). Springer. Haese RR, Hensen C, de Lange GJ (2006) Pore water geochemistry of eastern Mediterranean mud volcanoes: Implications for fluid transport and fluid origin. Mar Geol 225(1–4): 191–208. Henry P, LePichon X, Lallemant S, Lance S, Martin JB, Foucher JP, FialaMedioni A, Rostek F, Guilhaumou N, Pranal V, Castrec M (1996) Fluid flow in and around a mud volcano field seaward of the Barbados accretionary wedge: Results from Manon cruise. J Geophysical Research-Solid Earth 101: 20297–20323. Hensen C, Nuzzo M, Hornibrook E, Pinheiro LM, Bock B, Magalhaes VH, Bruckmann W (2007) Sources of mud volcano fluids in the Gulf of Cadiz - indications for hydrothermal imprint. Geochim Et Cosmochim Acta 71: 1232–1248.

Mud Volcanoes Jørgensen BB, Boetius A (2007) Feast and famine microbial life in the deep-sea bed. Nat Rev Microbiol 5(10): 770–781. Joye SB, MacDonald IR, Montoya JP, Peccini M (2005) Geophysical and geochemical signatures of Gulf of Mexico seafloor brines. Biogeosci Discuss, 2(3): 295–309. Judd AG, Hovland M, Dimitrov LI, Gil SG, Jukes V (2002) The geological methane budget at Continental Margins and its influence on climate change. Geofluids, 2(2): 109–126. Kaul N, Foucher JP, Heesemann M (2006) Estimating mud expulsion rates from temperature measurements on Hakon Mosby Mud Volcano, SW Barents Sea. Mar Geol 229: 1–14. Kopf A, Klaeschen D, Mascle J (2001) Extreme efficiency of mud volcanism in dewatering accretionary prisms. Earth Planet Sci Lett, 189(3–4): 295–313. Kopf AJ (2002) Significance of mud volcanism. Rev Geophys 40(2): B-1–B-49. Kopf AJ (2003) Global methane emission through mud volcanoes and its past and present impact on the Earth’s climate. Int J Earth Sci 92(5): 806–816. Lance S, Henry P, Le Pichon X, Lallemant S, Chamley H, Rostek F, Faugeres JC, Gonthier E, Olu K (1998) Submersible study of mud volcanoes seaward of the Barbados accretionary wedge: sedimentology, structure and rheology. Mar Geol 145(3–4): 255–292. Linke P, Suess E, Torres M, Martens V, Rugh WD, Ziebis W, Kulm LD (1994) In situ measurement of fluid flow from cold seeps at active continental margins. Deep-Sea Res I 41(4): 721–739. Linke P, Wallmann K, Suess E, Hensen C, Rehder G (2005) In situ benthic fluxes from an intermittently active mud volcano at the Costa Rica convergent margin. Earth Planet Sci Lett 235: 79–95. Lo¨sekann T, Knittel K, Nadalig T, Fuchs B, Niemann H, Boetius A, Amann R (2007) Diversity and abundance of aerobic and anaerobic methane oxidizers at the haakon mosby mud volcano, Barents Sea. Appl Environ Microbiol 73(10): 3348–3362. Mazzini A, Svensen H, Akhmanov GG, Aloisi G, Planke S, Malthe-Sorenssen A, Istadi B (2007) Triggering and dynamic evolution of the LUSI mud volcano, Indonesia. Earth Planet Sci Lett 261(3–4): 375–388. Mellors R, Kilb D, Aliyev A, Gasanov A, Yetirmishli G (2007) Correlations between earthquakes and large mud volcano eruptions. Journal of Geophysical Research-Solid Earth 112(B4): B04304. Milkov AV (2000) Worldwide distribution of submarine mud volcanoes and associated gas hydrates. Mar Geol 167(1–2): 29–42. Milkov AV, Sassen R, Apanasovich TV, Dadashev FG (2003) Global gas flux from mud volcanoes:

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A significant source of fossil methane in the atmosphere and the ocean. Geophys Res Lett 30(2): 1037. Mukhtarov AS, Kadirov FA, Guliyev IS, Feyzullayev A, Lerche I (2003) Temperature evolution in the Lokbatan mud volcano crater (Azerbaijan) after the eruption of 25 October 2001. Energy Explor Exploit 21(3) 187–207. Murton BJ, BiggsJ (2003) Numerical modelling of mud volcanoes and their flows using constraints from the Gulf of Cadiz. Mar Geol 195(1–4): 223–236. Niemann H, Duarte J, Hensen C, Omoregie E, Magalhaes V, Elvert M, Pinheiro LM, Kopf A, Boetius A (2006a) Microbial methane turnover at mud volcanoes of the Gulf of Cadiz. Geochim Cosmochim Acta 70(21): 5336–5355. Niemann H, Lo¨sekann T, de Beer D, Elvert M, Nadalig T, Knittel K, Amann R, Sauter EJ, Schlu¨ter M, Klages M, Foucher JP, Boetius A (2006b) Novel microbial communities of the Haakon Mosby mud volcano and their role as a methane sink. Nature 443: 854–858. Olu K, Lance S, Sibuet M, Henry P, FialaMedioni A, Dinet A (1997) Cold seep communities as indicators of fluid expulsion patterns through mud volcanoes seaward of the Barbados accretionary prism. Deep-Sea Res Part I-Oceanogr Res Pap 44(5): 811–841. Omoregie EO, Niemann H, Mastalerz V, de Lange G, Stadnitskaia A, Mascle J, Foucher JP, Boetius A (in review) Microbial methane oxidation and sulfate reduction at cold seeps of the deep Eastern Mediterranean Sea. Mar Geol accepted. Planke S, Svensen H, Hovland M, Banks DA, Jamtveit B (2003) Mud and fluid migration in active mud volcanoes in Azerbaijan. Geo-Mar Lett 23(3–4): 258–268. Sauter EJ, Muyakshin SI, Charlou JL, Schluter M, Boetius A, Jerosch K, Damm E, Foucher JP, Klages M (2006) Methane discharge from a deepsea submarine mud volcano into the upper water column by gas hydrate-coated methane bubbles. Earth Planet Sci Lett 243(3–4): 354–365. Schmincke HU (2006) Volcanism. Springer. Somoza L, Diaz-del-Rio V, Leon R, Ivanov M, Fernandez-Puga MC, Gardner JM, HernandezMolina FJ, Pinheiro LM, Rodero J, Lobato A, Maestro A, Vazquez JT, Medialdea T, FernandezSalas LM (2003) Seabed morphology and hydrocarbon seepage in the Gulf of Cadiz mud volcano area: Acoustic imagery, multibeam and ultra-high resolution seismic data. Mar Geol 195(1–4): 153–176. Stewart SA, Davies RJ (2006) Structure and emplacement of mud volcano systems in the South Caspian Basin. Am Assoc Pet Geol Bull 90(5): 771–786. Treude T, Boetius A, Knittel K, Wallmann K, Jorgensen BB (2003) Anaerobic oxidation of

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methane above gas hydrates at Hydrate Ridge, NE Pacific Ocean. Mar Ecol Prog Ser 264: 1–14. Vogt PR, Cherkashev A, Ginsburg GD, Ivanov GI, Crane K, Lein AY, Sundvor E, Pimenov NV, Egorov A (1997) Haakon Mosby mud volcano: A warm methane seep with seafloor hydrates and chemosynthesisbased Ecosystem in late Quantemary Slide Valley, Bear Island Fan, Barents Sea passive margin. EOS

Transactions of the American Geophysical Union Supplement 78(17): 187–189. Wallmann K, Linke P, Suess E, Bohrmann G, Sahling H, Schlu¨ter M, Da¨hlmann A, Lammers S, Greinert J, von Mirbach N (1997) Quantifying fluid flow, solute mixing, and biogeochemical turnover at cold vents of the eastern Aleutian subduction zone. Geochim Cosmochim Acta 61(24): 5209–5219.

14 Abiogenic Hydrocarbon Production at the GeosphereBiosphere Interface via Serpentinization Reactions G. Proskurowski Woods Hole Oceanographic Institution, Department of Marine Chemistry and Geochemistry, Woods Hole, MA, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 216 2 Serpentinization Leading to Reducing Conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 219 3 Reduction of Oxidized Carbon Species to Methane and Higher Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 220 4 Determining Abiotic Versus Biotic Hydrocarbon Sources . . . . . . . . . . . . . . . . . . . . . . . . . . . 223 5 Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 225 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 227

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_14, # Springer-Verlag Berlin Heidelberg, 2010

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Abiogenic Hydrocarbon Production in the Shallow Crust

Abstract: The production of hydrocarbons via mechanisms not associated with biological processes has far reaching implications to the fields of petroleum geochemistry, astrobiology, and the study of early life and life in extreme environments. Despite an intense focus on discovering abiotic hydrocarbon sources in natural settings, only a handful of sites convincingly suggest that abiotic organic synthesis occurs within the geosphere. Although experimental studies in aqueous settings clearly demonstrate the potential for abiotic synthesis, the scope of abiotic hydrocarbon production in natural settings has yet to be defined. As theoretical and experimental studies indicate that abiotic hydrocarbon synthesis is maximized in highly reducing environments, it is not surprising that the strongest evidence for abiotic organic synthesis in natural settings is associated with the alteration (serpentinization) of reduced mantle rocks. The crux of this topic is that currently there is no foolproof approach to distinguishing abiotic versus biotic organic synthesis. Thus, it is especially important to be cognizant of the possibilities and limitations of abiotic hydrocarbon production when considering a deep subsurface biosphere where the organic matter may be synthesized by both abiotic and biotic processes.

1

Introduction

The potential to abiotically produce hydrocarbons in natural settings, and in particular deepsea hydrothermal vent environments, has long piqued the interests of both scientists and the general public. The public’s curiosity primarily arises from the seemingly science fiction notion that simple inorganic constituents can self-assemble to form organic compounds that may represent prebiotic molecules from which life emerged or an energy source available to microbes and internal combustion engines (Gold, 1999). Recently, the economic pressures of a fixed petroleum supply responding to rapidly expanding global demand have renewed public interest in the potential for an undiscovered, inexhaustible abiotic source of petroleum. However, the public’s pining for an additional source of petroleum has yet to be supported by credible scientific evidence. And while there has been substantial scientific discussion surrounding the role of abiotic hydrocarbon production in natural settings to the origin of life (Holm and Andresson, 1998; Macleod et al., 1994; Martin et al., 2008; Russell and Hall, 2006), this debate remains largely theoretical. To date, the most concrete scientific implication of abiotic synthesis of methane and higher hydrocarbons in geological settings is the supply of metabolic energy and fixed carbon to biological communities of the sub-seafloor, seafloor, and overlying water column (Karl, 1995; Kelley et al., 2002; Kniemeyer et al., 2007). Here I discuss abiotic hydrocarbon production in serpentinizing crustal environments that highlight the transfer of biologically accessible carbon from the abiotic geosphere to the deep biosphere. Emphasis is on the likely mechanisms of abiotic hydrocarbon formation, and the current capability to determine the ultimate source of hydrocarbons by geochemical means. The source of hydrocarbon gases in geologic environments can be broadly classified into three sub-categories: ‘‘biogenic,’’ ‘‘thermogenic,’’ and ‘‘abiotic’’ (Schoell, 1988). Biogenic hydrocarbons are formed as a direct result of metabolic and biosynthetic processes (e.g., methanogenesis); thermogenic compounds are the result of the thermal maturation and degradation of living or diagenetically altered biomass (e.g., hydrothermal alteration of organic-rich sediments, ‘‘cracking’’ of kerogen). It is important to note that both biogenic and thermogenic hydrocarbon production require the presence of active or preformed biological material, and thus the ultimate source of carbon can be deemed ‘‘biotic.’’ On the

Abiogenic Hydrocarbon Production in the Shallow Crust

14

other hand, abiotic hydrocarbons are formed from inorganic starting materials by purely chemical processes that are totally independent of biological activity (e.g., Fischer-Tropsch type reactions). The primary difficulty of studying abiotic hydrocarbons in geologic settings is that natural samples are commonly generated by more than one mechanism, and are typically dominated by thermogenic and biogenic products in near-surface environments. Furthermore, the geochemical separation of biotic and abiotic sources is often ambiguous. Thus, biologically sourced hydrocarbons must be considered the default source until multiple lines of evidence suggest an abiotic source. Marine sediments serve as a reservoir for an extensive suite of organic matter (amino acids, carbohydrates, lignins, lipids, and uncharacterized organic matter) reflective of the biochemistry of terrestrial and marine organisms (Burdige, 2006). Where hydrothermal circulation intersects sedimentary environments the input of heated seawater results in the thermal decomposition of organic matter by mechanisms similar to the standard model of oil and natural gas production, but on much more rapid timescales (Seewald, 2003; Seewald et al., 1990). Hydrothermal vent fluids from organic-rich sedimentary environments such as Guaymas Basin (Gulf of California) and Middle Valley (Juan de Fuca Ridge) contain millimoles per kilogram concentrations of methane, micromoles per kilogram concentrations of C2–C4 alkanes and simple aromatic hydrocarbons, and detectable amounts of longerchained hydrocarbons, polycyclic aromatic hydrocarbons, and fatty acids (Cruse and Seewald, 2006; Simoneit et al., 1992, 1988). The lack of surface sediments at hydrothermal sites does not entirely preclude thermogenically produced hydrocarbons, as buried sediments (Proskurowski et al., 2004; You et al., 1994), or subsurface microbial communities could potentially provide a biological source of organic matter. However, unsedimented sites do provide a first-order filter that allows for an examination of abiotic hydrocarbons with minimal obfuscation by thermogenic inputs. For this reason the discussion is focused primarily on unsedimented, ultramafic-hosted hydrothermal systems, where the geological setting provides favorable chemical conditions for abiotic hydrocarbon synthesis while limiting the possibility for thermogenic production. Abiotic production of hydrocarbons in geologic settings can be generally described as the reduction of inorganic carbon (primarily CO2) by H2 to form CH4 and longer-chained linear hydrocarbons. This reaction is largely dependent on the redox conditions imparted by fluidrock interactions, primarily determined by the amount of H2 produced. The reduced ferrous iron, Fe(II), in olivine, pyroxene, and pyrrhotite minerals of basalt and ultramafic rocks provides an ideal reductant to convert H2O to H2 during alteration of the host rock during hydrothermal circulation. While the alteration of basalt can yield low concentrations of H2, the alteration of ultramafic rocks (serpentinization) liberates up to three orders of magnitude more H2 (McCollom and Seewald, 2007) (> Table 1). Ultramafic rocks, originating as either mantle peridotites or deep crustal plutonic rocks, can be emplaced in the shallow crust by tectonic processes. Seafloor exposures of ultramafic rocks occur along fracture zones, along passive margins, at the walls of rift valleys, and at mid-ocean ridge spreading centers (Fru¨hGreen et al., 2004). Ultramafic rocks are particularly abundant in slow- to ultra-slow spreading ridge environments where tectonic processes and a low magma supply lead to extensive faulting and uplift (Dick et al., 2003). Continental emplacements of oceanic ultramafic rocks primarily occur in obducted ophiolite sequences or in orogenic features accessed by alpine exposures or by deeply penetrating boreholes. Active hydrothermal and terrestrial seep sites located in ultramafic rocks present a rare opportunity to examine in-situ serpentinization and subsequent abiotic hydrocarbon

217

350

368

21
N EPRa

Endeavoura

317

352

Logatcheva

Guaymas

365

Rainbowa

3.2

0.4

1.7

12.0

16.0

14.19

13.26

9.22

5.43

4.20

3.69

52.00

3.40

0.07

2.10

2.50

1.55

1.55

1.07

1.07

1.29

1.31

1.84

1.21

1.44




78,000

n.m.

42

n.m.

1,097

1,390

1,470

970

970

1,080

1,140

1,700

1,150

1,220

1

n.m.

n.m.

n.m.

n.m.

n.m.

12

40

4

2

6

14

21

5

6,980

n.m.

26

n.m.

48

120

120

73

80

93

99

160

100

100

C3H8 (nmol kg−1)

1,100

n.m.

40

n.m.

n.m.

6

23

11

10

17

12

47

14

10

n-C4H10 (nmol kg−1)

410

n.m.

n.m.

n.m.

n.m.

5

5

14

10

1

5

4

9

9

i-C4H10 (nmol kg−1)

Chapter 12, Vol. 1, Part 3) are equivalent to those occurring in the deep subsurface ultramafic environments. Thus, the topic of abiotic hydrocarbon synthesis in serpentinizing systems has important implications for a subsurface microbial biosphere. The following discussion will summarize the potential mechanisms for abiotic hydrocarbon synthesis and the methods for diagnosing abiotic versus biotic production within the context of natural systems where hydrocarbons can be both a metabolic product of, and substrate for, microbial life.

2

Serpentinization Leading to Reducing Conditions

The reduction of oxidized inorganic carbon (primarily CO2, but also CO, and HCO3) to hydrocarbons in shallow crustal settings through geochemical reactions or thermodynamic equilibrium is largely governed by the reducing power of the fluid, i.e., the concentration of H2 and H2S. In fluids circulating through basaltic crust the reducing potential of the fluid is tempered by moderate H2 and H2S concentrations that are essentially fixed by reactions between seawater and the relatively oxidizing mineral assemblages containing plagioclase (Seyfried and Ding, 1995). In fluids circulating through ultramafic crust, highly reducing conditions result through the production of H2 associated with the oxidation of ferrous Fe-bearing minerals (e.g., olivine, pyroxene, and pyrrotite) to magnetite during

219

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Abiogenic Hydrocarbon Production in the Shallow Crust

serpentinization. The serpentinization of olivine, the dominant mineral in ultramafic rocks, can be described in the following reaction: 6½ðMg1:5 Fe0:5 ÞSiO4  þ 7H2 O ! 3½ðMg3 Si2 O5 ðOHÞ4  þ Fe3 O4 þ H2 olivine

water

serpentine

magnetitie

ð1Þ

hydrogen

Olivine is a solid solution of Mg- (forsterite) and Fe- (fayalite) olivine endmembers, and as such, the above depiction is just one of numerous reactions that can describe the hydration of olivine. Here the reactant is an iron-rich olivine (Fo75), and the reaction ignores other products (e.g., brucite, talc) that are possible with more Mg-rich olivines (Bach et al., 2006; Fru¨h-Green et al., 2004; Janecky and Seyfried, 1986). In general serpentinization reactions are irreversible, exothermic (2.2 kJ g1 H2O reacted), and lead to a rock volume expansion of up to 50% (Fyfe and Lonsdale, 1981; O’Hanley, 1992). However, generalities rarely apply when examining the thermodynamics and kinetics of serpentinization. For example, the temperature at which serpentinization occurs (determined by 18O thermometry) varies widely, from 100
C in subduction zone settings to 450
C in select mid-ocean ridge settings (Fru¨h-Green et al., 2004). Geochemical models are challenged by variable reaction conditions (e.g., temperature, pH, water to rock ratios, rock compositions), the complexity of product mineral phases including Fe-substituted minerals (e.g., Fe-serpentine, Fe-brucite), and evolving reaction mechanisms as fluids move through progressively altered rock. Although notable modeling efforts have made impressive strides in simulating the complexity of serpentinization (Allen and Seyfried, 2003, 2004; Alt and Shanks, 2003; Foustoukos et al., 2008; Janecky and Seyfried, 1986; Seyfried et al., 2007; Wetzel and Shock, 2000), model predictions continue to overestimate the H2 concentrations resulting from serpentinization. A recent theoretical and experimental examination of serpentinization on a mineral assemblage representative of the ultramafic basement at Lost City at 200
C and 500 bar by Seyfried et al. (2007), resulted in model predicted H2 concentrations of >300 mmol kg1, and experimental yields of 75 mmol kg1. Seyfried et al. (2007) hypothesizes that this discrepancy is due to H2 production associated not only with magnetite formation (as suggested in reaction 1), but also with Fe-bearing alteration phases for which limited thermodynamic data exist. This important conclusion suggests significant H2 production can occur during the initial stages of serpentinization, without the formation of magnetite (a late-stage product), in effect expanding the role of serpentinization as a source of H2 to fluids circulating through ultramafic crust. The three ultramafic-hosted hydrothermal sites Logatchev, Rainbow, and Lost City all have maximum H2 concentrations of 16 mmol kg1 (see > Table 1, > Fig. 1), about five times less H2 than produced in the experiments of Seyfried et al. (2007). This inconsistency is likely explained by slight differences between the environmental conditions and the experimental setup (e.g., temperature, water to rock ratios, rock composition) and becomes less pronounced when considering the possibility for further reaction of H2 in the production of hydrocarbons.

3

Reduction of Oxidized Carbon Species to Methane and Higher Hydrocarbons

While serpentinization reactions alone do not produce hydrocarbons, the reducing conditions resulting from H2 formation enhance the potential for the abiotic synthesis of organic matter. Hydrocarbon production in natural settings will only result if favorable thermodynamic

Abiogenic Hydrocarbon Production in the Shallow Crust

14

. Figure 1 CH4 and H2 concentrations from a variety of hydrothermal vent sites. Lost City, Logatchev, and Rainbow sites are the only non-sediment-hosted sites with high concentrations of CH4 and H2. Middle Valley is a sedimented site on the Juan de Fuca Ridge; the high H2 concentrations at 9
N EPR were sampled directly after an eruptive event and reflect the alteration of fresh basalt.

conditions exist and the time frame of reaction is sufficiently long as to not be kinetically inhibited. Perhaps the simplest abiotic hydrocarbon formation mechanism is the equilibrium reaction between CO2 and CH4: CO2 þ 4H2 , CH4 þ 2H2 O

ð2Þ

As illustrated in > Fig. 2, the thermodynamics of this reaction favors CH4 production at lower temperatures and/or higher H2 concentrations, and the dominance of CO2 at typical basalt-hosted hydrothermal conditions (T > 300
C, H2 of 100 mmol kg1). However, this equilibrium reaction is slow to proceed under hydrothermal conditions, as illustrated by the extremely limited amount of kinetic data from uncatalyzed experimental studies at T < 450
C (Horita, 2001). When catalyzed by transition metals (notably, Ni, Cr, Pt, Pd) the irreversible form of this reaction, known as the Sabatier process, proceeds in hydrothermal experiments (Berndt et al., 1996; Foustoukos et al., 2004; Horita and Berndt, 1999) and has been invoked in natural settings (Fiebig et al., 2007). In the geochemical literature the Sabatier process is often misidentified and subsumed into the more expansive Fischer-Tropsch type (FTT) reaction that produces longer-chained hydrocarbons in addition to methane: CO2aq þ ½2 þ ðm=2nÞH2 ! ð1=nÞCn Hm þ 2H2 O

ð3Þ

Because CO2 is the dominant oxidized carbon source in hydrothermal environments the more general term Fischer-Tropsch type is used in place of the more exacting Fischer-Tropsch synthesis (FTS), which refers specifically to the reduction of CO. Although CH4 and higher hydrocarbons have been synthesized by FTS in the gas phase from CO for nearly 100 years

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Abiogenic Hydrocarbon Production in the Shallow Crust

. Figure 2 Thermodynamic equilibrium calculations for the reaction between CO2 and CH4 at hydrothermal temperature and H2 conditions. CH4 formation (the reverse of the reaction as written) is favorable only at temperatures less than 350
C for the highest H2 conditions, and less than 250
C for typical H2 conditions Figure courtesy of T. McCollom.

(Anderson, 1984), only recently were FTT reactions shown to proceed, albeit with low yields, under aqueous hydrothermal conditions (temperatures 175–400
C), with dissolved CO2 as the carbon source (Foustoukos and Seyfried, 2004; McCollom and Seewald, 2006). The use of the term ‘‘FTT’’ indicates the assumption that the FTS reaction is preceded by CO production from CO2 via the water–gas shift reaction: CO2 þ H2 , CO þ H2 O

ð4Þ

The FTS proceeds as CO binds to the surface of a metal catalyst, is reduced to carbide and then methylene (–CH2–), and then polymerizes with other methylene groups to form alkyl chains that eventually terminate with a methyl group (–CH3) or surface bound H. This polymerization model for chain growth leads to a characteristic suite of linear alkyl chains, dominated by methane and the longer alkanes, with lesser abundances of the alkenes, alkynes, and trace amounts of oxidized hydrocarbons such as alkenones, alkanols, and alkanoic acids. The relative abundance of FTS products commonly exhibits a log-linear decrease in abundance with increasing carbon number, known as the Anderson-Schulz-Flory distribution (Anderson, 1984). As will be discussed in the next section, this distribution is characteristic of, but not diagnostic of, FTT. It has become commonplace to invoke Fischer-Tropsch type reactions when discussing the abiotic origin for hydrocarbons in geologic settings. This presumption of mechanism is largely based on the compatibility of the FTT reaction with the environmental settings. Indeed, the intersection of mantle rocks and water at submarine ultramafic-hosted systems (Charlou et al., 2002; Proskurowski et al., 2008), springs issuing from ophiolites (Abrajano et al., 1988), deeply penetrating boreholes and mines (Sherwood Lollar et al., 2002, 2006), and gabbroic fluid inclusions (Kelley, 1996; Kelley and Fru¨h-Green, 1999) provides the moderate temperature ( Table 1 and > Fig. 1). While longer-chained hydrocarbons (primarily alkanes) concentrations have been reported from sedimented sites (Cruse and Seewald, 2006), and from Lost City (Proskurowski et al., 2008) and Rainbow (Charlou et al., 2002), only a handful of data exists for unsedimented sites (McCollom and Seewald, 2007). The concentration of CH4 relative to the C2+ hydrocarbon concentration has been proposed as a tool to distinguish microbial, thermogenic, and abiotic sources (Horita and Berndt, 1999). A long held view of C1/C2+ ratios is that a thermogenic C1/C2+ signal is less than 100, while microbial C1/C2+ values range from 2,000 to 13,000 (Simoneit et al., 1988). The C1/C2+ ratios derived from experimental hydrothermal FTT studies range broadly from 3,000 (Horita and Berndt, 1999), which is not surprising considering the highly varied experimental catalytic and reaction conditions. The C1/C2+ values from Lost City (1,000) and Rainbow (2,000) are greater than typical thermogenic values, and fall within the broad range of experimental FTT results. However, mixing of microbial and thermogenic sources would also result in the observed C1/C2+ values, thus limiting the diagnostic utility of this approach. The stable hydrogen and carbon isotopic composition of hydrocarbons provide additional parameters by which sources can be discriminated. A widely employed geochemical tool to determine the source of CH4 is a plot of the dD-CH4 versus d13C-CH4 (Schoell, 1980; Whiticar, 1990), where microbially produced CH4 is highly depleted in D and 13C, thermogenic CH4 is moderately depleted in D and 13C, and abiotic CH4 is relatively enriched in D and 13 C. However, this isotope-isotope source plot has major shortcomings: (1) absolute isotope values do not describe isotope fractionation relative to an initial carbon source, (2) hightemperature and high-pressure conditions greatly reduce isotope fractionation associated with microbial processes (Takai et al., 2008), and (3) carbon-limited environments may result in isotopic compositions that are not reflective of the process-oriented fractionation (Bradley et al., 2009; Proskurowski et al., 2008). While the use of plots describing source–product fractionation (aDH2O–CH4 vs a13CDIC–CH4) (Sherwood Lollar et al., 2008) may eliminate variations in the source isotopic composition (e.g., organic matter vs magmatic CO2), in many cases the isotopic fractionations involved in abiotic and biotic processes may be remarkably similar. Thus, it is recommended that CH4 source plots be used only as a comparative tool rather than as a diagnostic one. The isotopic composition of CH4 and higher hydrocarbons does contain information suggestive of origin, however there is debate over the proper interpretation. The d13C-CH4 from the three ultramafic-hosted hydrothermal sites Lost City, Rainbow, and Logatchev is 9 to 16‰, enriched in 13C relative to all other hydrothermal sites. However, experimental FTT work by McCollom and Seewald (2006) show a 36‰ fractionation between the source carbon and CH4, suggesting that CH4 at these sites should be more depleted in 13C if it is abiotic in origin. Proskurowski et al. (2008) argue that in carbon-limited environments the d13C-CH4 approaches the d13C of the source carbon, which at Lost City is 9‰.

Abiogenic Hydrocarbon Production in the Shallow Crust

14

The isotopic trends in dD and d13C of linear alkanes with increasing chain length are a promising, but not definitive, means to distinguish thermogenically and abiotically produced hydrocarbons. A typical thermogenic trend, shown in > Fig. 3, is defined by isotopic enrichments in 13C with increasing chain length (d13C1 < d13C2 < d13C3 < d13C4). In contrast, abiotically produced hydrocarbons from Lost City (Proskurowski et al., 2008), Kidd Creek (Sherwood Lollar et al., 2002, 2008), and experimental work (Hu et al., 1998; McCollom and Seewald, 2006; Taran et al., 2007) exhibit a carbon isotopic trend that is slightly depleted or relatively flat with increasing chain length. While this ‘‘reverse thermogenic trend’’ in 13C isotopic composition appears to reflect abiotic synthesis, it may not be unique to abiotic processes, as similar patterns have been observed in thermogenic gases from pyrolyzed lignite (Du et al., 2003). Furthermore, standard thermogenic 13C isotopic trends have been measured from abiotic hydrocarbons produced experimentally (Fu et al., 2007). The hydrogen isotopic trend for thermogenic alkanes is typically characterized by isotopic enrichments in D with increasing chain length (dD–C1 < dD–C2 < dD–C3 < dD–C4). However, the hydrogen isotopic trends for abiotic volatile hydrocarbons are less well defined, as fewer measurements have been made. C1–C3 alkanes from Lost City exhibit a relatively flat hydrogen isotopic trend with small and variable changes in dD with increasing chain lengths ( Fig. 1 gives an overview of the utilization of coal and its conversation. Pyrolysis is the heating of fossil raw material (e.g., coal, lignite, peat, wood) producing gases, liquids, and solids at temperatures higher than 300
C in an oxygen-free atmosphere. General decomposition products are the pyrolysis gas or town gas (composed mainly of hydrogen, carbon monoxide, carbon dioxide, methane, nitrogen, and C2–C5 hydrocarbons), the liquid products (tar oil, crude benzene, and water),

Coking Processes and Manufactured Gas Plants

15

. Figure 1 Coal utilization and conversation.

and coke as a solid residue. Industrial sites especially those that are primarily produced town gas are often referred to as Manufactured Gas Plants or ‘‘MGPs’’. The general process of is divided into several steps: 1. 2. 3. 4. 5.

Coal is heated in a retort. Gas passes through a condenser to remove tar. Other impurities (e.g., ammonia) are eliminated by a scrubber. Hydrogen sulfide is removed by passing the gas over trays containing moist ferric oxide. The resulting gas is finally stored in a gasholder.

The actual process parameters depend on the target product and they influence the composition of the by-products. Using coal as feedstock at low carbonization temperature (below 700
C), one can obtain fine coke and large quantities of liquid and gaseous products, while high-temperature treatment (900–1,300
C) produces coke for blast furnaces (Crelling et al., 2006).

3

Characterization of Tar Oil

About 15  106 t/a of coal tar is still produced worldwide primarily as a by-product of coke processing (Collin and Ho¨ke, 2005). The composition as well as the properties of tar oil is mainly influenced by the raw material and process parameters, e.g., treatment temperature (> Table 1). For example, low-temperature tar contains a much higher amount of phenols compared with high-temperature tar. Also lignite, peat, or wood tars have a high proportion of phenols, while the carbon content is much lower and the water content higher. With increasing geological age of the raw material, the content of phenols and solid paraffins decreases while the ‘‘aromaticity’’ increases. High density indicates a high degree of aromatization. Coke oven tar with a density of ca. 1.2 g/cm3 contains the highest fraction of aromatic hydrocarbons (naphthalene content is around 10%).

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Coking Processes and Manufactured Gas Plants

> Table 2 gives an overview of a fraction distillation analysis of a coal tar. This indicates that tar oils are multicomponent mixtures. According to Collin and Ho¨ke, 2005, it contains more than 10,000 different constituents. Most substances (up to 85%) are PAHs, while others are phenols (already mentioned), mono aromatic compounds, and NSO-heterocyclic compounds. The tar oil content of selected substances and their physico-chemical properties are given in > Table 3.

. Table 1 Properties of various tars (Colin and Ho¨ke, 2005) Coal tars Cokeoven tars

Lowtemperature Gasification tars tars

Lignite tars

Peat tars

Wood tars

Density at 20
C [g/cm3]

1.14–1.25 0.96–1.05

1.05–1.14

0.95– 1.05

0.94– 0.98

1.08–1.20

Carbon [%]

90–93

83–85

84–86

81–85

78–82

60–65

Hydrogen [%]

5–6

8–9.5

6–8

9–11

8–10

6–8

Naphthalene [%]

5–15

0–2

2–4

0–1

0–1

0–0.5

Phenols [%]

0.5–5

10–45

15–20

5–30

5–25

20–40

Bases [%]

0.2–2

0.5–2

0.5–4

0.1–15

0.5–3

traces–0.5

Solid paraffins [%]

0–traces

3–15

0–5

8–20

10–30

traces

Coking residues (ISO 6615:1983) [%]

10–40

5–15

10–20

4–9

4–10

10–18

Toluene insolubles [%]

2–20

0.5–10

3–10

0.2–1.5

2–8

traces–12

Ash (800
C) [%]

Fig. 2).

3

Performing a ‘‘Dirty Job’’: Oil Pollution in Shipping Operations

Marine fuel oil: Every modern ship uses different types of fuel oil for propulsion dependent on the type of their engine. During and mainly after the Second World War, the global merchant fleets were transformed from ‘‘steamers’’ using coal and steam boilers to fuel oil propelled ships. The advantages of this measure were obvious - lack of a need of stokers reduced the ship’s crew sizes and the danger of fires and explosions in coalbunker was eliminated.

. Figure 2 Causes and magnitude of marine oil pollution, 1973–1993.

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Two major types of engines were constructed: the combustion (‘‘diesel’’) engines are built for reliability and low fuel consumption. These engines are classified according to their operating speed. Slow speed engines are crosshead two-stroke engines with a speed up to 300 rpm are commonly used in freighters and tankers, whereas medium speed engines are limited to battleships and cruise liners. Their trunk or opposed piston four stroke engines are able to operate at speeds up to 900 rpm. This enables a higher velocity up to 35 knots, the equivalent of 65 km/h. A second principle of ship propulsion is the naval gas turbine. Like the four stroke diesel engine, a turbine operates at high rotation speeds and thus is commonly used in war ships and increasingly in cruise liners. Gas turbines use combusted air and highly volatile fuels to create a stream of hot gases that are subsequently transformed in kinetic energy when led through the turbine’s blades. Turbines allow higher speeds than diesel engines and are smaller in size. Additionally they can also be used to power an electric engine or to generate a water jet for propelling the vessel (King, 1956). However, the drawback with turbine engines is that they consume much more fuel compared to other engine types. Dependant on their propulsion principle, ships use different types of fuel: turbine engines must use light fuel oils such as kerosene or marine diesel oil, medium-speed four stroke diesel engines can additionally use marine gas oil. Slow speed two stroke engines can use the whole spectrum of fuels including heavy fuel oils such as Bunker C (King, 1956). These oils used for propulsion are one of the main sources of oil pollution in shipping: they can get spilled during refuelling (‘‘bunkering’’), during maintenance work and in case of a grounding or accident (Hampton et al., 2003; Huijer, 2005). Since their engines consume up to 14 tons of fuel oil per hour, large container ships such as the state-of-the-art containership ‘‘Emma Maersk’’ have bunker capacities of up to 10,000 metric tons, the equivalent of a small oil tanker. Because of this fact, even cargo ship averages can lead to large-scale oil spills and serious pollutions. Lubricant oil: In parallel to fuel oils, all types of ships require a large variety of lubricant oils. Since marine diesel engines can weigh up to 2,500 tons and reach sizes of a small house, they consume large amounts of lubricants for pistons, valves, gears, shafts and other engine parts. These lubricant oils in contrast to fuel oils are mostly synthetic oils containing paraffin oils, wax and additives. Altogether, these mixtures of hydrocarbons are hardly biodegradable. Both fuel and lubricant oils as well as other run-off such as anti-fouling substances, cement, chemicals, detergents, iron oxides, paints and solvents, together with the crew’s domestic waste water, are collected in the lowest compartment of the ship; the bilge (King, 1956). Bilge waters and illegal discharge: Bilge waters result from seawater being washed into the ship in rough seas and rainy weather. This water maintains a variety of substances and is gathered in the bilge. Although bilge water therefore is a noxious, corrosive aqueous mixture, which needs to be disposed in adequate ways in ports, seaborne waste management makes ship operations safer and easier for the crew. However, since bilge water disposal is also very expensive, it is mostly processed onboard (Hampton et al., 2003). The main problem of bilge water is the oil content, which can be reduced by oil-water separators (OWS) commonly combined with pollution control systems. After oil separation and filtration of particles, the remaining bilge water can be pumped overboard. However, in many cases, the onboard oil separator capacities do not match the occurring amounts of bilge water. Therefore, OWS systems and pollution control systems can be bypassed by a so-called ‘‘magic pipe,’’ which conveys untreated bilge water into the sea. By falsification of records, bilge water, toxic sludge and other liquid waste can be disposed without payment (Corbet and Koehler, 2003; Hampton et al., 2003). This kind of waste disposal is forbidden according to the MARPOL convention

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and persecuted in many countries (IMO, 2002a). However, the persecution of this kind of pollution requires airborne control and chemical analysis of water samples, which cannot be provided in all countries. The amount of pollution due to ‘‘magic pipes’’ and other forms of illegal dumping of liquid and solid waste can only be estimated. Tank washing: Another serious type of ship-borne pollution is a product of tanker operations, arising from the cleaning of tanks after oil has been carried in them (Brockis, 1967). After unloading, oil residues remain on the walls of the oil tanker’s compartments. These residues lead to a major problem: an oil tanker is build for maximum manoeuvrability and seaworthiness when fully loaded. This problem occurs in all cargo ships, which are designed to carry a maximum of cargo in relation to the ship’s size. Empty cargo ships therefore loose pitch stability and risk capsizing in rough sea or storms. Furthermore, rudders and propeller could not be completely immersed in the sea, what leads to reduced manoeuvrability and inefficient propulsion (King, 1956). To stabilize or ‘‘trim’’ a ship, all ships need to store a certain amount of water in specific ballast tanks. While ballast water in cargo ship is not toxic (it might even carry biological material), ballast water of oil tankers poses a major source of oil contamination. Tankers, which do not possess separate ballast tanks, typically fill several compartments of the hull (‘‘oil tanks’’) with seawater. This leads to contamination of the ballast water with the oil residues that remain on the compartments’ sides and bottoms. The other compartments of the tanker were also cleaned with seawater and detergents until the 1960s. Both the ballast water and the emulsions resulting from tank washing were pumped overboard, leading to grave pollution (Brockis, 1967). Alternative tank washing strategies were developed in the 1960s and 1970s, mostly due to the losses of oil resulting from this procedure. The first procedure developed was the so-called ‘‘load on top’’ – method (LOT). The tanks were washed as described before using hot seawater and surfactants. After cleaning of the tanks, the oily washings and dirty ballast water are pumped into a so-called ‘‘slop tank.’’ Some of the clean tanks are refilled with fresh seawater. During the voyage, oil and water in the slop tank are separated. The lower, aqueous phase is then pumped overboard. When arriving at its destination, fresh crude oil is loaded into the clean tanks and on top of the oily residues in the slop tank. This procedure can be repeated very often until finally a cleaning of the slop tanks will be necessary. Load on top cleaning saves up to 800 tons of oil on large tankers and thus increased the profit of the ship’s owners, but the main beneficiary was the environment (Brockis, 1967). An even better improvement was the ‘‘crude oil washing’’ (COW) – method developed in the 1970s. This procedure is based on the observation, that crude oil is most effectively washed by oil. Minor technical modifications on oil tankers enabled to wash tanks with crude or other types of oil. Oil is sprayed on the residues clinging to the walls of the tanks and dissolving it completely. After the voyage of the tanker, the residues are unloaded with the rest of the oil (Brockis, 1967; IMO, 2002b). However, the use of this method is limited to tankers transporting the same type of oil. Tanker transporting heavy crude or fuel oil can not use COW when transporting light oils, diesel, kerosene or gasoline (Brockis, 1967; IMO, 2002b). The light oils get contaminated with flocks of heavy fuel oil, which block tubes and pipes when pumping or burning the diesel/kerosene in power plants or engines. Nevertheless COW has reduced the problem of slops and oil pollution due to tank washing almost completely (Brockis, 1967). Crude oil washing is mandatory to all oil tankers larger than 20,000 dwt since 1978 according to the MARPOL convention (> Fig. 3) (IMO, 2002a, b). Modern oil tankers avoid all these problems by technical means: especially larger ships include a separate system for ballast water enabling trimming the ship without oil

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. Figure 3 Procedures in tank washing, ‘‘load on top’’ (LOT) and ‘‘crude oil washing’’ (COW).

250 Shipping-Related Accidental and Deliberate Release into the Environment

Shipping-Related Accidental and Deliberate Release into the Environment

16

contamination. Furthermore, the new generation of tankers also include separate tanks for collecting residues resulting from crude oil washing (Hampton et al., 2003). But what are the sources of oil pollution in the new millennium? Statistics on the oil pollution due to tanker shipping show that there is an occurrence of small scale oil spills smaller than 7 tons of oil that are the result of bunkering and discharging (Huijer, 2005). The reason for this could be many-fold: A broken valve or pipe, poor maintenance or human error. All in all, these oil pollutions can be avoided by proper safety standards and adequate maintenance, which might sometimes be sacrificed in a business under the high pressure of maximising profits. Therefore, pollution hot spots are the tanker terminals that are often located in the oil producing countries which have poor public resources to ensure safety standards and enforce legislation concerning environmental protection. Last but not least, oil transportation is a dangerous business involving large machinery and applying huge mechanical forces. A 350 m VLCC supertanker weighs approximately 500,000 tons, the equivalent of 50 Eiffel towers. Moving such a mass involves enormous kinetic energy, which can easily destroy tubes, pipes or machinery in docks or bunkering terminals. High performance pumps installed on the ships or in the terminals are likely able to pump 10,000,000 kg of oil into or out of a tanker per hour. Even minor malfunctions therefore can lead to significant volumes of oil being spilt. Although safety regulations and the use of smaller and well-designed tankers including state-of-the-art technologies and materials will decrease the pollution risks from oil transportation, not even total abandonment of using crude oil as a source of energy and raw material might prevent oil pollution. A ‘‘gift’’ from the past: Ship wrecks and ‘‘Environmental timebombs’’: Oil pollution in the sea has many sources: they could be of natural origin, such as algal blooms or marine ‘‘seeps’’ – cracks in oil fields on the ocean’s floor. They could be of anthropogenic origin, such as run-off from industry and cities, transported by rivers or from shipping as has been presented above. However, statistics concerning oil spills caused by shipping clearly focus on present day and the recent past. Since large-scale oil pollution became a problem in the early 1960s, the earliest records of this environmental problem are dated back to this time (Huijer, 2005). In fact, a far bigger threat still rests below the ocean’s surfaces. In 2001, mysterious oil pollution occurred in the Ulithi Lagoon at the island of Yap in the Pacific Ocean. 91 tons of oil contaminated the lagoon and prevented fishing (the only source of income for the 600 members of the island community) (Christie, 2002). An investigation quickly found the source of the pollution and showed that Yap was in fact on the brink of an ecological catastrophe. Being scheduled to assist in an American landing operation in the Gulf of Leyte on 19th October 1944, the USS ‘‘Mississinewa,’’ an American T2 oil tanker, fuelled with 1,500 tons of kerosene, 1,000 tons of diesel and 10,000 tons of fuel oil, was anchored in the Ulithi Lagoon. In the morning of October 20th, an explosion struck the 169 m long Mississinewa. Within a few minutes, the highly explosive cargo of the tanker caught fire and exploded. The Mississinewa capsized and sank within minutes with the loss of 63 sailors. In fact, the oil tanker was hit by a Kaiten, a Japanese manned torpedo, designed for Kamikaze operations. The ship’s wreck remained in a depth of 40 m in Ulithi Lagoon until finally in 2001 the Mississinewa’s hull breached due to corrosion in the warm seawaters of the Pacific Ocean (Christie, 2002; Prevatt, 2003; Mair, 2007). While most of the volatile diesel and kerosene were either spilled, burned or evaporated, the majority of the fuel oil remained in the 22 intact tanks of the wreck. In an attempt to avoid further destruction in Ulithi Lagoon, a salvage diving team of the US Navy recovered the remaining 7,600 tons of fuel oil (Prevatt, 2003). The

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. Figure 4 Oil released from the wreck of the USS Mississinewa in 2001 (left); sinking of the Mississinewa in 1943 (right) (source: NAVSEA).

Mississinewa incident drew attention to an environmental problem of unknown size which had been forgotten for more than half a millennium: Ship wrecks, which remained in the sea after the Second World War on all marine battlefields of the earth (> Fig. 4) (Christie, 2002; Prevatt, 2003). Naval warfare of the Second World War and its ‘‘heritage’’ in the Second Millennium: The beginning of World War II marked the start of a catastrophe for merchant shipping. The early military success of the German Third Reich quickly provided the German Navy with access to the Atlantic Ocean. The only remaining opposition to German oppression in Europe in 1939 was therefore posed by the United Kingdom. Since an invasion of the British islands was not possible for the Third Reich, a campaign to interrupt the support of food supply, goods and energy sources was undertaken by attempting an embargo (Rohwer, 1999 & 2007). Due to the Treaty of Versailles, the German Navy was clearly outnumbered and outgunned by the Royal Navy in that time. Subsequently, major efforts to disrupt merchant shipping were concentrated on submarine warfare. Due to the availability of experienced ship builders and submarine captains from World War I, the expertise and effectiveness of the ‘‘Submarine Weapon’’ was established in a short time. From 1938 to 1939, large numbers of newly build submarines went into service in North Sea and the Western Approaches southwest of Ireland (Costello et al., 1977). Because of the low numbers of small and fast battleships such as destroyers or corvettes and the relatively old English merchant fleet, high numbers of cargo ships and tankers in the vicinity of the British islands were sunk in 1939 and 1940, leading to a difficult supply situation in the United Kingdom. The reaction to this situation was the formation of convoys of several dozens of merchant ships, guarded by battleships and destroyers (Costello et al., 1977). Although a convoy allows the efficient use of the protecting war ships, the speed of the whole convoy must equal the speed of the slowest merchant ship. Most of these ships however used coal burning steam engines. Early convoys therefore were travelling at 6 to 8 knots (11 to 13 km/h) what made them highly susceptible to submarine attacks (Rohwer, 2007). A newly developed tactic in submarine warfare, the ‘‘wolf pack’’ tactic, involving several submarines simultaneously attacking the same convoy, had devastating effects on these early

Shipping-Related Accidental and Deliberate Release into the Environment

16

convoys (Costello et al., 1977; Rohwer, 2007). Using this strategy, a total cut-off of sea trade on the North Atlantic could have been reached by 300 submarines organised in ‘‘wolf packs’’ (Rohwer, 2007). In 1939 and 1940, 667 merchant ships – more 22% of the British merchant fleet - were sunk in the Western Approaches and around the British Islands (Costello et al., 1977; Rohwer, 1999). The lack of transportation capacities and – even worse – experienced sailors (more than 10,000 sailors died) worsened the supply situation in the United Kingdom and lead to an intervention of the United States: 50 destroyers and merchant ships were lent to the United Kingdom (Rohwer, 2007). Following the attack on Pearl Harbour, the USA was forced into Second World War in December 1941. As a support for their British allies, the largest ship building operation of all times began. The idea of this program was to replace ships destroyed by German submarines, battleships and airplanes faster than they could be sunk. This was the beginning of a battle of material in the Atlantic Ocean (> Fig. 5) (Rohwer, 2007). The shipbuilding program involved a standard design of ship, modular standardized ship hulls combined with standardized components and equipment. New wielding techniques increased the speed of construction by 600% (From approximately 260 to 40 days. The fastest construction time for the so-called ‘‘Liberty’’ ships was 108 h). These ships were equipped with steam turbines or diesel engines consuming oil instead of coal (Spyrou, 2006; Marine Log, 2008). However, qualitative and quantitative improvements of the German U-boats, submarine tactics and crew experience lead to a stalemate situation. In 1942, the submarines in North Atlantic were able to sink a total of 1150 ships (Costello, 1977). This disastrous loss of life and material led to further improvements in the Battle for the North Atlantic. Improved standardized freighters (‘‘Victory’’-Ships) and tankers (‘‘T2-Tankers’’ such as the Mississinewa) were constructed. Using oil-fuelled diesel and turbine engines, these ships reached a velocity of 12 to 17 knots (28–31 km/h). This increased speed made them and thus the convoys as a whole faster and less susceptible to submarine attack (a submerged standard German Type VIIC/41-submarine could only reach a speed of 7 knots) (Costello et al., 1977; Rohwer, 2007). By the middle of 1943, the submarine battle in the Atlantic was lost. The development of sound detection systems (‘‘ASDIC,’’ better known as SONAR), RADAR and the deciphering of the German ‘‘Enigma’’

. Figure 5 Scenes of the submarine warfare from 1939 to 1945. Left picture: Allied tanker torpedoed. World War II torpedoes were partially equipped with magnetic ignitors, making them explode below a ship’s keel. Hulls therefore were breaching and ships often broke apart in two parts (US National Archives and Record Administration; ARC Identifier: 520607). Right picture: German Type IX-C submarine (source: Australian War Memorial, AWM).

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. Figure 6 Locations convoy routes and Allied merchant ships as well as German submarines sunk in the initial phase of World War II (1939–1940) and at the climax of the convoy battles (1942–1943) (source: Costello et al.). (Costello et al., 1977; Rohwer, 2007)

radio code made further submarine operations futile. Furthermore, the loss of experienced crews (replacement crews consisted of 17 year old sailors commanded by 20 year old captains in 1944) lead to grave losses and the end of the convoy battles (> Fig. 6) (Rohwer, 2007). While submarine warfare in Atlantic was ending, it was merely beginning in the Pacific Ocean. American submarines, airplanes and battleships started to win superiority over the Japanese Navy and merchant fleet. Japan, an industrial power but with poor natural resources, possessed a large merchant navy mostly constructed in the 1920s and 1930s. During the Second World War, the Japanese tankers and freighters were needed not only to supply the Japanese industry with resources, but also to provide supplies to the islands invaded during the late 1930s. Furthermore, the Japanese battle fleet needed to be supported with fuel and supplies. Therefore, the destruction of cargo ships was a method of choice in the defeat of the Japanese Empire (Roscoe, 1950). The naval battles of Second World War led to tremendous losses of lives and ships. 30,000 sailors on allied merchant ships died and 28,000 out of 40,000 submarine crewmen did not return. Due to the nature of the submarine war in the Atlantic, only 175 allied warships were sunk, but in contrast a horrendous number of 3,550 merchant ships were lost (Rohwer, 2007). In the pacific theatre, 1,703 freighters and tankers were destroyed (Roscoe, 1950). The example of the ‘‘Mississinewa’’ shows the dangerous potential of the 4,200 shipwrecks, which remain on the seafloor after World War II (> Fig. 7). Thankfully most of the ships sunk in the deeper region of the Atlantic have suffered severe hull breaches when sinking to the ocean’s bed and thus released most of their cargo. In many

Shipping-Related Accidental and Deliberate Release into the Environment

16

. Figure 7 The destruction of shipwrecks caused by currents. Dependent on the position of a wreck in a river or a tidal current, undercutting can take place in different positions of the shipwreck, such as bow and stern or in midship position. Continuous undercutting leads to the formation of ‘‘trenches’’ which destabilise the wreck. Corrosion and stress to the ships structure eventually cause hull damage.

cases, the tankers on the Atlantic transported light oils such as gasoline, kerosene or diesel to fuel trucks, airplanes and tanks. These ships often went ablaze as described by eyewitnesses (Costello et al., 1977; Prevatt, 2003; Rohwer, 2007). The majority of the hydrocarbons therefore burned, evaporated or were biodegraded quickly. A major threat is posed by freighters and tankers sunk in shallow waters, such as lagoons in the Pacific Ocean, i.e., in the Philippines, Guadalcanal etc. There are additional threats to the marine ecosystems from wrecks found in the shallower waters surrounding the British Isles. Records of the ships sunk in 1939 and 1940 show an accumulation of shipwrecks in Bay of Biscay, Shetland Islands, Dogger Bank, English Channel and the Western Approaches. Corrosion of these wrecks is one problem, while the loss of structural integrity due to currents is another. Although most of the merchant ships had robust wielded hulls made of steel, corrosion is slowly degrading the ship’s bottoms, decks and bulkheads. The corrosion should be faster in warm waters such as the shallow Pacific but events comparable to the Mississinewa incident are only a matter of time in the colder European waters. The destruction of shipwrecks due to undercutting is a further problem known for tearing apart, displacing or deforming shipwrecks. This phenomenon can mostly be observed in rivers. Due to differences in the speed of currents occurring at different parts of a wreck, sediment can be removed below the bow and stern of the wreck. The shipwreck therefore can be bent and – in the worst case – break apart (Dehling, 2001).

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Research Needs

The Second World War – sometimes also referred to as the worst catastrophe in the history of mankind – provides humanity with many problems, even 75 years after it ended. The submarine ‘‘heritage’’ of the naval warfare will challenge salvage specialists for several more decades. The ecological time-bombs resulting from this period may not be detected at all. Some might be deactivated by simple means, and others will require expensive salvage operations. At the end of the day, they must be a reminder from the past to future generations, never to let the spectre of war to be seen again.

References Brockis GJ (1967) Preventing oil pollution of the sea. Helgoland Mar Res 16: 296–305. Christie M (2002) World War Two Wrecks Haunt Pacific with Oil Spills. Australia: Reuters News Service, November 4. Corbett JJ, Koehler HW (2003) Updated emissions from ocean shipping. J Geophys Res 108: 233–259. Costello J, Hughes T, Collins M (1977) The Battle of the Atlantic, London: Book Club Associates. Cowan E (1968) Oil and Water – the Torrey Canyon Disaster. Philadelphia: Lippincott. Crude Oil Washing (2002) International Maritime Organization (IMO). www.imo.org. Dehling T (2001) Wracksuche. Bundesamt fu¨r Seeschifffahrt und Hydrographie. www.bsh.de. January 11. Hampton S, Kelley PR, Carter HR (2003) Tank vessel operations, seabirds, and chronic oil pollution in California. Mar Ornithol 31: 29–34. Heideloff C (2004) ISL Shipping Statistics and Market Review. Institute of Shipping Economics and Logistics. www.isl.org. Huijer K (2005) Trends in Oil Spills from Tanker Ships 1995–2004. London: International Tanker Owners Pollution Federation (ITOPF). International Maritime Organization (IMO) (2002a) MARPOL Convention. Protocols and Annexes. www. imo.org. International Maritime Organization (IMO) (2002b) Crude Oil Washing. www.imo.org. King GAB (1956) Tanker practice. The Construction, Operation and Maintenance of Tankers. Wokingham: Maritime Press. ¨ lverschLink PM (2000) Gefa¨hrdungspotentiale von O mutzungen durch Schiffshavarien in der Nordsee dargestellt am Beispiel der Amoco Cadiz und der Pallas, Diploma thesis, Geographisches Institut, Christian-Albrechts-Universita¨t zu Kiel, Germany.

Mair M (2007) Oil, Fire and Fate: The Sinking of the USS Mississinewa AO-59 in WWII by Japan’s Secret Weapon. Santa Ana, CA: Seven Locks Press. Marine Log (2008) The Liberty Ship and the T-2 Tanker (1941) Ships of the Century. www.marinelog.com. MARPOL Convention (2002) Protocols and Annexes. International Maritime Organization (IMO). www. imo.org. National Research Council (NRC) (1985) Oil in the Sea, Inputs, Fates and Effects. Washington, DC: National Academy Press. Prevatt J (2003) USS Mississinewa (AO 59) oil offloading operations, Ulithi Atoll, February 2003. Naval Sea Systems Command (NAVSEA). Marine Environmental Engineering Technology Symposium, August 20–22. Rohwer J (1999) Axis Submarine Successes of World War Two (German, Italian and Japanese Submarine Successes, 1939–1945). London: Greenhill Books. Rohwer J (2007) Chronology of the War at Sea 1939– 1945. Online Edition, www.wlb-stuttgart.de/seek rieg/chronik.htm, June 11. Roscoe T (1950) United States Submarine Operations in World War II. Annapolis: US Naval Institution. Spyrou AG (2006) From T-2 to Supertanker: Development of the Oil Tanker, 1940–2000. Lin-coln, NE: iUniverse, Inc. Tanker practice (1956) The Construction, Operation and Maintenance of Tankers. King GAB. Wokingham: Maritime Press. The Liberty Ship and the T-2 Tanker (1941) Ships of the Century. Marine Log. www.marinelog.com. April 8, 2008. USS Mississinewa (AO-59). Wikipedia, The Free Encyclopedia. http://en.wikipedia.org/wiki/USS_Missis sinewa_(AO-59).

17 Oil Tanker Sludges and Slops C. Gertler1 . M. M. Yakimov2 . M. C. Malpass3 . P. N. Golyshin1 1 School of Biological Sciences, Bangor University, Bangor, Gwynedd, UK [email protected] [email protected] 2 Institute for Coastal Marine Environment, Messina, Italy [email protected] 3 School of Environment and Natural Resources, Bangor University, Bangor, Gwynedd, UK 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 258 2 Oil Tanker Sludges and Slops . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 261 3 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 262

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Abstract: Despite international agreements, economic interests, and increasing environmental protection awareness, oil transportation in the third millennium is a dangerous business causing many cases of pollution. It is a paradigm for the negative effects of globalization, as has been proven by a product tanker in 2006. Following an odyssey, which took her from Europe to Africa, a gasoline tanker entered the port of Abidjian, Ivory Coast on 19 August 2006. Her presence and activities in the port of Abidjian claimed 16 lives and caused more than 40,000 cases of poisoning.

1

Introduction

The voyage of the Panama-registered tanker vessel began from Gibraltar on 26 June 2006. While visiting the port of Amsterdam 1 week later, the tanker was chartered by a Dutch trading company, which attempted to process waste and tanker slops by the port reception facilities. During the inspection of the tanker slops, an immense smell was reported by the waste disposal company’s staff. As a result, the port-based company refused to dispose the waste (Bernard et al., 2006; Franoz and Gesret, 2008). An analysis of the tanker slops in the aftermath of the Abidjan catastrophe showed that the gasoline tanker indeed was used as a ‘‘swimming refinery’’ (Persson, 2006). On her last voyage prior to the visit in Amsterdam, the vessel was transporting naphtha, a gasoline precursor containing significant amounts of mercaptans – sulfur-containing hydrocarbons. As these sulfur-containing hydrocarbons should be removed to meet the specifications for gasoline in certain European countries, a chemical process called ‘‘Merox’’ was conducted. By adding specific catalysts, water, and sodium hydroxide to the gasoline, mercaptans are oxidized to disulfides. The products of Merox are upgraded gasoline and sludge containing sulfuric components (Wikipedia, 2006). In the case of the tanker vessel, this chemical process failed due to the addition of too much sodium hydroxide and resulted in a highly toxic sludge containing mercaptans and hydrogen sulfide (> Fig. 1). After failing to dispose this toxic waste, the charterer attempted to arrange the disposal of the toxic slops using a Dutch waste disposal company, which quoted 500,000€ for this service (Bernard et al., 2006; Persson, 2006). Unwilling to pay this fee, the gasoline tank was ordered to continue its voyage to the Estonian port of Paldiski, where it was bunkering gasoline bound for Lagos, Nigeria. The gasoline was loaded on top of the toxic sludge. After unloading the gasoline in Lagos, the charterer attempted once again to dispose the toxic waste, but decided to keep the slops in the ship because the Nigerian company seemed to intend to refine it and sell it as fuel. As a result, the tanker vessel was called to the port of Abidjan, Ivory Coast. Although Abidjan was a modern port with

. Figure 1 Chemical reactions of the Merox (mercaptan oxidation) process.

Oil Tanker Sludges and Slops

17

adequate port reception facilities, it did not provide a slop disposal service. This work was performed by a newly incorporated waste disposal company. This company offered the disposal of the slops at a price of US$ 35 per cubic meter, 20 times less than the price offered in the port of Amsterdam (Bernard et al., 2006). The tanker discharged the slops and continued its voyage back to Paldiski (> Fig. 2). A few weeks later, citizens of Abidjan started to complain about previously unknown symptoms of diseases following heavy rains. Some suffered from nosebleeds, vomiting, headaches, and rashes. Most of them described the water with a smell of rotten eggs or garlic that was running through the streets of Abidjan. More than 40,000 Ivorian citizens showed symptoms described above, 69 were taken to hospitals and 16 died (Bernard et al., 2006). Although the charterer claimed only to have handed regular tanker slops to the Abidjanbased waste disposal company, analysis showed that indeed the highly toxic sludge also was transferred to a barge and subsequently into tanker trucks (Bernard et al., 2006). The disposal company, obviously overstrained with the disposal of toxic waste, dumped 500,000 l of toxic waste at 14 locations in and around Abidjan, i.e., lakes, rivers, fields, or simply in roadside trenches (Bernard et al., 2006; Franoz and Gesret, 2008). Truck drivers claimed that they were told to ‘‘dispose dirty water’’ by their managers (Bernard et al., 2006; Franoz and Gesret, 2008). Following the heavy rains, the waste was slowly distributed over the whole rural and municipal area of Abidjan, a city with a population of three million.

. Figure 2 Route of the Panama-registered and Dutch-chartered tanker vessel involved in the 2006 Coˆte d’Ivoire toxic waste spill.

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In the aftermath, many top government officials resigned (Bernard et al., 2006). Two executives of the charter company offered assistance, but were arrested in Ivory Coast upon arrival. The charterer ultimately paid US$ 198 million for compensation and cleanup (Bernard et al., 2006; Franoz and Gesret 2008) (> Fig. 3). Although this incident was an extremely severe case of tanker slop dumping, it displays an alarming trend of global trade. Corrupt officials and administrations often are eager or forced into offering inadequate waste disposal for enterprises based in industrialized countries. In 1988, the government of a West African country offered waste disposal including radioactive and chemical waste for US$ 1.6 million down payment and 30 years of economic aid (Knauer et al., 2006). The deal was canceled after a public row and media opposition. In parallel to the illegal disposal of tanker slops in ports as presented above, tanker slops often are disposed directly into the sea.

. Figure 3 Sampling on a spill site in Abidjan (right) (From CEDRE).

Oil Tanker Sludges and Slops

2

17

Oil Tanker Sludges and Slops

Tanker slops are result of the need for tank washing when changing the type of cargo after a trip. Oil residues remain on the tank’s sides and bottom. Contamination of the refined hydrocarbons particularly occurs when changing from a crude oil to a refined product. In the first 10 decades of tanker shipping, tanks were routinely washed with seawater and detergents, the washings were combined in a tank until a phase separation occurred and the aqueous phase dumped overboard. This measure leads to immense oil pollution and therefore was outlawed (King, 1956). Two other measures – the Load On Top and the Crude Oil Washing (COW) – were implemented and have led to a drastic decrease in oil pollution (Hampton et al., 2006; Yvonnou, 2001). Both measures are presented in Essay 00527. COW reduces the amount of oil spilled during tanker operations to a minimum, since the oil tanks are washed with crude oil that dissolves the residues. But if COW has reduced this marine oil pollution to zero, why do tanker slops still occur and why do they pose a threat? As described in > Chapter 16, Vol. 1, Part 3, tanker slops are a result of the need to fill some oil-contaminated tanks with seawater as ballast in order to stabilize the ship at sea and prevent capsizing in heavy seas. The first measure to prevent this was tank washing and subsequent loading of cargo oil on to the washings. The amount of slops was reduced to 300–800 m3 on large tankers transporting more than 320,000 m3 of oil. In most cases, refineries accepted the slops and used them for refining purposes. Refineries, ship owners, and charterer benefited from this arrangement. However, the need for highly refined products finally stopped the use of this method in the 1980s (Hampton et al., 2006; Yvonnou, 2001). COW was implemented and reduced slops from 800 to 250 m3 on large vessels. To reduce tanker slops to zero, modern tanker designs include segregated ballast tanks (SBT). SBT tankers are highly advantageous, because they are considered ‘‘ecological’’ vessels that produce no slops. On the other hand, refineries receive a neat product from these tankers: oil not contaminated with seawater, chemicals, or oil residues. Oil and water cannot come into contact on such SBT tankers, and still the ship’s seaworthiness is maintained. Since furthermore the tanks are washed using crude oil, there should, in theory, be no further slops occurring. However, in the real world, two situations cause slops on all tanker vessels including SBT vessels. The first situation is the compulsory washing of tanks prior to maintenance being carried out on tanks, pipes, or valves either on board or in shipyards. The tank washings in theory should be collected, settled, and kept. In the best case scenario, these slops can be separated via a water or oil separator. Oil is retained and the water can be disposed of. In modern vessels the purity of disposed water furthermore is controlled by detectors fitted to the seawater exit valves. However, these control systems can be bypassed and the slops can be directly dumped into seawater (See > Chapter 16, Vol. 1, Part 3). The second situation that might produce tanker slops is due to a safety measure for heavy sea and stormy weather. The SBT in an SBT tanker’s hull are designed to maintain the seaworthiness of a tanker at regular conditions. In a 270,000 dead weight tons (dwt) tanker, the ballast tanks may hold 60,000–70,000 t of seawater. In storm conditions such as hurricanes, the amount of ballast water is increased to 100,000–150,000 t on the same vessel. The tanker has a deeper draught and is less susceptible to sheering forces due to wind or waves. Therefore, the forces affecting the 300–350-m long ship’s hull are reduced and thus the risks of damage or, in worst case, sinking. The additional ballast water, however, has to be transferred to some of

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the oil-polluted cargo tanks. This emergency measure leads back to the initial problem of tanker slops and raises the question what might happen to slop in the cargo tanks when it comes to dispose of the emergency ballast water (Hampton et al., 2006; Yvonnou, 2001). But even if an SBT tanker, applying COW, is operated in proper manner attempting to minimize pollution, slops might still occur. Firstly, certain oils contain high amounts of paraffin or wax. Like all oils transported in tanker vessels, these oils are heated in order to maintain a low viscosity of the product. Relatively low temperatures of the tank’s walls lead to the deposit of waxy residues, which cannot be reheated and remain in the tanks after unloading. Secondly, a tanker vessel needs a good stern trim and properly maintained COW pumps in order to reduce the slops. If the inclination of a tanker to the stern or bow is too severe, COW pumps cannot produce enough suction to perform at optimum efficiency or reach oil residues collected in distant parts of the tank (Hampton et al., 2006; Yvonnou, 2001). Furthermore, technical equipment such as pumps are constantly stressed and endangered by corrosion on the high seas, reducing their performance and leading to the production of tanker slops. As described above, tanker slops are not simply limited to crude oil carriers, but also occur in product tankers such as the vessel involved in the 2006 Coˆte d’Ivoire toxic waste spill. Considering the chemical reactions potentially conducted inside the ships’ cargo tanks during voyages, it can only be assumed that how many other cases of comparable toxic slops are produced. Following the 2006 incident, no further accidents of this kind have been reported. However, catalysts and caustic soda for the Merox process are freely available and can also be applied for upgrading liquid petroleum gas (LPG), which is frequently transported by specialized LPG tankers. Perhaps the 2006 Coˆte d’Ivoire incident has scared tanker owners from making further attempts of on-board processing of oils. Nevertheless, the opportunity and the economic benefit to perform it remain.

3

Research Needs

Unfortunately, the occurrence of slops in the operations of tankers is inevitable. A number of technical methods to avoid or minimize slops have been invented and are currently employed. For the slops that nevertheless occur, proper disposal is possible in most ports of call. However, as we have presented above, this method of disposal is expensive and the urgent need for it can be abused by waste disposal companies. This economic pressure subsequently leads to the temptation of illegally discharging or dumping slops. The only possibility of preventing dumping of oil slops is the routine surveillance of tanker routes and prosecution of identified polluters. Many industrialized countries have developed efficient systems of identifying and prosecuting ships suspected of dumping slops together with methods of assigning oil pollution to the individual vessel. For surveillance, highly specialized airplanes are used for regular reconnaissance flights on frequently used shipping routes. To patrol North and Baltic Sea the German Air Force uses four Fairchild-Dornier DO 228 LM airplanes equipped with side-scanning radar, IR/UV line scanner, LASER-fluoro-scanners, and microwave radiometers. Their equipment is able to identify the type of oil spilt and measures the thickness and extent of oil slicks while in flight. Ships causing pollution, therefore, can be easily identified and countermeasures coordinated. Typically, samples will be taken by coast guard ships and identified by automated gas chromatography. This method enables the authorities to receive a ‘‘fingerprint’’ – a qualitative analysis of the components of the oil spilled (Dahlmann, 2003; Wikipedia, 2008). A ship

Oil Tanker Sludges and Slops

17

spotted while dumping slops or discharging contaminated water will be apprehended by the coast guard and forced to call at the nearest port. In case of an oil slick floating on the ocean’s surface, sampling and subsequent analysis data enable authorities to compare the pollution to ballast water samples taken from suspicious ships (> Fig. 4). Reconnaissance flights are routinely performed over shipping lanes of the North and Baltic Seas. The HELCOM treaty, signed by states bordering the Baltic Sea, mandates the signatory countries to collaborate together and conduct regular monitoring of the Baltic, and thus gives rise to increased levels of surveillance. This treaty was signed by Denmark, Estonia, Finland, Germany, Latvia, Lithuania, Poland, Russia, and Sweden in order to prevent ship collisions, eutrophication, and oil pollution of the Baltic Sea. Aerial surveillance of the seas around European countries is a relatively simple task, as shipping routes are short, close to land, and traffic-dense. Furthermore, the industrialized European states have appropriate funds and large air forces/navies to undertake aerial surveillance duties. An equivalent system exists in Japan, the USA, and Canada, though shipping routes surrounding these countries are longer and traffic spread over large distances, aerial reconnaissance is more difficult. States with smaller funds or less strict environmental policies cannot offer this facility or prosecute the polluters. In some cases, corruption can be a major problem in the enforcement of environmental policies. But even if aerial surveillance is performed, airplanes and their detection systems rely on favorable weather conditions and sunlight. In stormy weather or at night, surveillance flights are difficult and inefficient. In combination with the large number of ‘‘black spots’’ on the map without appropriate surveillance, there are many opportunities for potential polluters to

. Figure 4 A Do 228 LM – reconnaissance airplane (ßBundeswehr/Cherin Hellmich).

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dump tanker slops or to discharge the toxic waste. A solution to the protection of the blue vastness of the ocean lies in the black infinity of space; in the last few decades, a significant number of satellites equipped with specific infrared (IR), microwave RADAR, and other cameras have been launched and are available for aerial reconnaissance. The satellite’s sensitive cameras are capable of performing a similar role to reconnaissance aircraft, are cost-effective, and can simultaneously survey much larger areas. Since they can even survey the shipping routes around countries, which cannot perform other measures of detecting oil pollution, satellite-based surveys may be a solution to this problem (Brekke and Solberg, 2004). However, aerial reconnaissance, oil analysis, and prosecution of illegal dumping only treats the symptoms rather than the cause. In the aftermath of most recent cases of oil pollution due to tanker slops or ballast water, the consequences have been similar. In general, ships were detained in a port until a trial for pollution has started. In many cases, the ship’s senior officers were sentenced to imprisonment, faced a financial penalty, or loss of licenses and thus their jobs. The ship’s charterers were sentenced to pay fines ranging from US$10,000 up to 500,000 (CEDRE, 2009). However, in an industry under the high pressures of transportation costs and delivery dates, the risk of paying such fines can be acceptable from an economic point of view. In order to limit oil pollution, the European Union (EU) implemented the directive 2000/59/EC on the basis of a ‘‘polluter pays principle’’: port reception facilities had to be provided by any harbor in the EU. The additional costs for this service were included in the harbor dues or charged as a separate fee. Ships that cause less pollution benefited from this system since reduced fees were charged for lower emissions. As a result, several harbor dues schemes including a free slop disposal service within the fees were set up by a number of port authorities. This free service was readily accepted and significantly reduced the pollution in areas such as the North Sea. However, in recent years, strong competition between the European port companies has occurred. In many cases, slop and waste disposal was performed by subcontractors rather than the port authorities and companies. Shipping companies, being offered several ports in similar locations, such as Antwerp, Esbjerg, Felixstowe, Rotterdam, Amsterdam, Bremerhaven, or Hamburg were able to call at the port charging the lowest harbor dues. The dues therefore had to be reduced to maintain the utilized capacities and thus the free disposal service had to be canceled in many cases. Furthermore, the slop disposal during berthing at a specific port was not mandatory according to the EU directive (Georgakellos, 2007). However, this example of ‘‘free’’ disposal service shows a potential solution to the problem of tanker slops. Effective prosecution of polluters and draconian penalties for illegal dumping may be good deterrents, but can only act after pollution has already occurred. It is a negative control mechanism, which is considered as an obligation. Moreover, the lack of a mandatory slop disposal at every port of call leaves the decision of disposal to the ship’s owner or charterer. Analysis of waste disposal figures for EU port reception facilities for 2000 and 2001 showed approximately 12% of liquid wastes generated in berthed ships that were actually disposed of at their reception facilities. This implies that the current system of waste disposal in the EU is either not strict enough or does not offer satisfactory incentives or deterrents for the ship’s charterers and owners in order to dispose slops safely. Illicit discharge or inadequate waste disposal in developing countries therefore cannot be prevented with the current system of harbor dues (Georgakellos, 2007). A mandatory, free disposal service, which is partly refunded by harbor dues, offers a compromise. Costs for the disposal thus should be shared by charterers, ship owners, and port management together with the consumers of the transported goods. Another important factor

Oil Tanker Sludges and Slops

17

is the global implementation of such disposal services. If mandatory disposal fees combined with mandatory disposal at every port are a constant factor in the accounts of all charterers and ship owners, there is no further interest in avoiding disposal or illegal dumping of slops. Eliminating the mechanism of the free market in environmental issues such as tanker slop disposal could be a very efficient method of significantly improving the situation of maritime oil pollution. It is nevertheless a long and hard path to follow, but considering the consequences of the 2006 Coˆte d’Ivoire incident, i.e., more than 40,000 victims and a clean-up costs of US$ 200,000,000 in a single case – it might be a path worth traveling on.

References Bernard F, Follorou J, Stroobants JP (2006) How Abidjan Became a Dump. Guardian Weekly 20 October 2006. Brekke C, Solberg AHS (2004) Oil spill detection by satellite remote sensing. Remote Sens Environ 95: 1–13. CEDRE (2009) Discharge at sea: Illicit discharge. Accessed on 16/1/09 from http://www.cedre.fr/uk/ discharge/ill_disch/cas.htm. Dahlmann G (2003) COSI – computerized oil spill identification. Bundesamt fuer Schifffahrt und Hydrographie, June 2003. Franoz B, Gesret S (2008) The Probo Koala. Information days of Cedre, 18 March 2008, INHES, Saint-DenisLa-Plaine (Paris). Georgakellos GA (2007) The use of the deposit–refund framework in port reception facilities charging systems. Marine Pollut Bull 54: 508–520. Hampton S, Kelley PR, Carter HR (2006) Tank vessel operations, seabirds, and chronic oil pollution in California. Mar Ornithol 31: 29–34. King GAB (1956) Tanker practice. The construction, operation and maintenance of tankers. Wokingham: Maritime Press.

Knauer S, Thielke T, Traufetter G (2006) Profits for Europe, industrial slop for Africa. Der Spiegel 38/2006. Persson M (2006) Petrochemie met de handboeken dicht. De Volkskrant 2 October 2006. Wikipedia contributors (2006) Coˆte d’Ivoire toxic waste spill [Internet]. Wikipedia, The Free Encyclopedia 2008 December 27, 21:03 UTC. Accessed on 13/1/09 from http://en.wikipedia.org/w/index.php? title = 2006_C%C3%B4te_d%27Ivoire_toxic_waste_ spill&oldid = 260376418. Wikipedia contributors. Dornier Do 228 LM. Wikipedia, The Free Encyclopedia 2008 Jul 14, 10:59 UTC. Accessed on 13/1/09 from http://de.wikipedia.org/ w/index.php?title = Dornier_Do_228_LM&oldid = 48367460. Wikipedia contributors. Merox [Internet]. Wikipedia, The Free Encyclopedia; 2008 December 27, 13:01 UTC. Accessed on 13/1/09 from http://en.wikipedia.org/ w/index.php?title = Merox&oldid = 260317814. Yvonnou LA (2001) Discharge at Sea – what about slops, Captain? 21 March 2001. Accessed on 16/1/09 from www.cedre.fr.

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Part 4

Environmental Chemistry

18 Chemistry of Volatile Organic Compounds in the Atmosphere R. Koppmann University of Wuppertal, Physics Department – Atmospheric Physics, Wuppertal, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270

2 2.1 2.2 2.3 2.4

Chemistry of VOCs in the Atmosphere . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 270 Degradation of Atmospheric Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 272 Degradation of Atmospheric Alkenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 Degradation of Atmospheric Aromatic VOCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276 Degradation of Atmospheric OVOCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 276

3

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 277

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Chemistry of Volatile Organic Compounds in the Atmosphere

Abstract: To put it simply, the cycle of VOCs in the atmosphere can be summarized as follows: VOCs are emitted from anthropogenic and natural sources mainly as gaseous, often non-polar compounds of high vapor pressure. Photochemical oxidation reactions involving OH and NO3 radicals, but also ozone and in some cases chlorine atoms, transform these compounds into mainly polar, water soluble compounds of low vapor pressure. These products are finally removed from the atmosphere by dry or wet deposition. At the very end of the reaction chains the final products are water vapor and carbon dioxide. While most of the VOCs themselves, especially at the relatively low concentrations are harmless, the products formed during the oxidation of VOCs in the atmosphere such as photo oxidants like ozone or PAN have a significant impact on air quality and can be harmful to human health.

1

Introduction

Every day, large quantities of volatile organic compounds (VOCs) are emitted into the atmosphere from both human activities and natural sources. The term ‘‘VOC’’ comprises a sheer limitless number of compounds. The major classes of compounds from anthropogenic sources are saturated hydrocarbons (alkanes), unsaturated hydrocarbons (alkenes and aromatic compounds such as benzene, toluene, and xylenes), and oxygenated compounds such as aldehydes, ketones, alcohols, esters, etc. Biogenic sources, mainly terrestrial vegetation, emits unsaturated compounds, preferably isoprene, but also monoterpenes and sesquiterpenes. Almost in the same amount also aldehydes and alcohols are emitted from vegetation. > Table 1 gives an overview of sources and amounts of emitted VOC. VOCs are present in the global atmosphere at mixing ratios of some 10 ppbv (parts per billion, 10
9, or nmol/mol) down to some ppt (parts per trillion, 10
12, or pmol/mol). Despite these low concentrations, VOCs have profound effects in the atmosphere. They are the ‘‘fuel’’ which keeps atmospheric photochemistry running. Therefore, their sources, sinks, residence times, and (photo)chemical reaction pathways were subject of research in the last three decades and still are an important objective of current research. They influence photochemistry on a local, regional, and even global scale. Some compounds have a potential impact on climate, both due to their properties as greenhouse gases, and also through their ability to form secondary organic aerosol (SOA) particles on oxidation.

2

Chemistry of VOCs in the Atmosphere

The degradation of VOC in the atmosphere is initiated mainly by the reaction with the OH radical, which itself is formed by the reaction of an O(1D) atom with a water molecule following the photolysis of an ozone molecule at wavelengths of l < 340 nm.
O3 þ hn ! O2 þO 1 D 


O 1 D þ H2 O ! 2 OH

The OH radical has an average global abundance of about 1  106 radicals/cm3. Despite this low concentration, OH is the most important cleansing agent in the troposphere. In addition to OH, also ozone and the nitrate radical (NO3) contribute to the degradation of VOCs, mainly unsaturated compounds.

Chemistry of Volatile Organic Compounds in the Atmosphere

18

. Table 1 Annual global emission rates of VOC from different anthropogenic and natural sources. All data are given in Tg C per year (Table taken from Williams J, Koppmann R (2007). Chapter 1 in Koppmann R (ed.) Volatile Organic Compounds in the Atmosphere. Oxford: Blackwell Publishing.) Emission rate

Uncertainty range

Alkanes

28

15–60

Alkenes

12

5–25

Aromatic compounds

20

10–30

Fossil fuel use

Biomass burning Alkanes

15

7–30

Alkenes

20

10–30

5

2–10

Aromatic compounds Terrestrial plants Isoprene

460

200–1,800

Sum of monoterpenes

140

50–400

Sum of other VOC

580

150–2,400

Oceans Alkanes

1

0–2

Alkenes

6

3–12

Sum of anthropogenic emissions Alkanes

44

Alkenes

38

Aromatic compounds

25

Terrestrial plants

1,180

Total

1,290

The nitrate radical, NO3, is formed by the reaction NO2 þ O3 ! NO3 þ O3 and is only present in the nighttime atmosphere due to its fast removal by photolysis during daytime. In addition, NO3 also reacts with NO2, forming dinitrogen pentoxide in a reversible equilibrium reaction NO3 þ NO2 þ M $ N2 O5 þ M Therefore, the mixing ratios of NO3 at night are low, typically between a few ppt and a few 100 ppt. In contrast to OH and NO3 radicals, ozone is ubiquitous in the troposphere at mixing ratios of some 10 ppb in clean environments and peak values of >100 ppb during photochemical smog episodes. Under specific circumstances such as in the marine boundary layer, also the reaction with chlorine atoms may play a role in the degradation of VOCs. However, if chlorine atoms

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are available, their concentrations are very low, typically a few 1,000 atoms per cm3. Therefore, regarding atmospheric chemistry in the troposphere on a global scale, chlorine atoms are of minor importance and usually not considered in global atmospheric chemistry models. The loss rate of any VOC is determined by the concentrations of the VOCs and the corresponding oxidants and can be calculated by d½VOC ¼
kOxidant ½Oxidant½VOC dt where [X] is the concentration of the oxidant and the compound of interest and k the rate coefficient (i.e., the velocity) of the corresponding reaction. The atmospheric residence times of VOCs are determined by the concentration of the oxidant and the corresponding rate coefficient. tVOC ¼

1 kOxidant  ½Oxidant

Average lifetimes of VOCs in the atmosphere range from about 8 years for methane to a few minutes for some sesquiterpenes. > Table 2 gives an overview of the average atmospheric lifetimes of some compound groups and a number of selected VOC. A schematic of the oxidation of atmospheric VOCs is shown in > Fig. 1. In the following the atmospheric degradation of different VOC groups is discussed in more detail.

2.1

Degradation of Atmospheric Alkanes

The reaction of saturated hydrocarbons with OH radicals proceeds by the abstraction of an H atom and the formation of a water molecule and an alkyl radical. The typical reaction pathway is shown for methane as an example: 

OH þ CH4 !  CH3 þ H2 O

The resulting alkyl radical, a methyl radical in this case, reacts very fast in a three-bodyreaction with O2 to form a methyl peroxy radical (M denotes any other molecule, because of their abundance typically N2 or O2 molecules): 

CH3 þ O2 þ M ! CH3 O2 þ M

In the presence of NO (>10 ppt) oxygen is abstracted to form a methoxy radical: CH3 O2 þ NO ! CH3 O þ NO2 The NO2 molecule formed in this reaction is photolysed leading to a ground state oxygen atom, which reacts almost immediately with an oxygen molecule to form ozone: NO2 þ hn ! NO þ  O 3 P 





O 3 P þ O2 þ M ! O3 þ M

Chemistry of Volatile Organic Compounds in the Atmosphere

18

. Table 2 Overview of average tropospheric lifetimes of VOC compound groups and some selected VOC as examples. Lifetimes are given for an average OH concentration of 6105 cm
3 and an average ozone concentration of 71011 cm
3 (about 30 ppb). (Table taken from Williams J, Koppmann R (2007) Chapter 1 in Koppmann R (ed.) Volatile Organic Compounds in the Atmosphere. Oxford: Blackwell Publishing.) Compound

Average lifetime

Alkanes

months – days

Ethane

2.5 months

Propane

2.5 weeks

n-Pentane

4 days

Alkenes

days – hours

Ethene

1.5 days

Propene

11 h

1-Butene

10 h

Cyclic compounds

days – hours

Cyclopentane

4 days

Methylcyclohexane

2 days

Cyclohexane

3h

Aromatic compounds

weeks – hours

Benzene

2 weeks

Toluene

2 days

1,3,5-Trimethylbenzene

7.5 h

Biogenic compounds

hours – minutes

Isoprene

3h

a-Pinene

4h

Limonene

30 min

In this way, the oxidation of VOCs in the presence of NO is the main process responsible for the production of ozone in the troposphere. Especially during smog events in summer, urban air masses contain high concentrations of both VOC and NO. Intensive photochemistry during the transport of these air masses leads to the extremely high ozone levels which are often observed in rural areas downwind from urban sources. The methoxy radical reacts with O2 to form formaldehyde as the first stable product from this reaction chain. CH3 O þ O2 ! CH2 O þ HO2 In very clean environments the methoxy radical can also react with other peroxy radicals, (i.e., HO2 or organic peroxy radicals). Reaction with HO2 results in the formation of more stable methyl hydroperoxides: CH3 O þ HO2 ! CH3 OOH þ O2

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. Figure 1 Schematic of the oxidation of atmospheric VOCs into the first stable carbonyl products in the presence of NOx, and the formation of ozone. (Figure taken from Le Bras G (2002) Gas Phase Reactions, Encyclopedia of Atmospheric Science. In JR Holton (ed.). Amsterdam: Elsevier, pp. 352–359).

In the same way higher molecular weight alkanes are attacked by OH radicals via hydrogen abstraction, as illustrated in the following using R as a symbol for an organic rest such as CH3CH2
in case of ethane, for instance. RH þ  OH !  R þ H2 O For longer chained alkanes with different types of hydrogen-carbon bonds, reaction rates for the H abstraction are decreasing with the number of hydrogen atoms attached to the same carbon atom. Therefore, the abstraction is favored as follows: =CH– > –CH2– > –CH3. The resulting alkyl radical reacts fast with O2 to form an alkyl peroxy radical: 

R þ O2 þ M ! RO2 þ M

In the presence of NO (>10–30 ppt) oxygen is abstracted to form an alkoxy radical: RO2 þ NO þ M ! RO þ NO2 Again, NO2 is produced leading to tropospheric ozone formation, whereas the alkoxy radical, depending on the molecule, can thermally decompose, isomerize or react with molecular oxygen. Breaking of the carbon chain leads to the formation of a stable aldehyde and a smaller peroxy radical, which again can react as described above. Isomerization occurs by internal hydrogen abstraction and eventually leads to a hydroxycarbonyl molecule. The reaction with O2 leads to the formation of a stable ketone via the abstraction of a hydrogen atom, which is attached to the same carbon atom as the oxygen radical. Furthermore, alkyl peroxy radicals can also attach to an NO or an NO2 molecule. In the case of NO this leads to the formation of relatively stable organic nitrates, whereas with NO2 peroxynitrates are produced:

Chemistry of Volatile Organic Compounds in the Atmosphere

18

RO2 þ NO þ M ! RONO2 þ M RO2 þ NO2 þ M ! RO2 NO2 þ M The yields of the organic nitrates increases with the chain lengths of the alkanes. The most important peroxynitrate is the peroxyacetylnitrate or PAN (CH3C(O)O2NO2). Since this compound is not directly emitted by human activities, it is an excellent measure for photochemical processing of an air mass and an important component of photochemical smog. PAN and organic nitrates are important in relation to long-range transport in the atmosphere, because they act as reservoirs for reactive nitrogen as they are fairly stable at low temperatures. In very clean environments the peroxy radicals can also react with other peroxy radicals, (i.e., HO2 or organic peroxy radicals). Reaction with HO2 results in the formation of more stable organic peroxides: RO2 þ HO2 ! ROOH The combination with another peroxy radical is either a sink for the radicals and produces alcohols, ketones and organic acids or leads to alkoxy radicals, which react as described above: RO2 þ RO2 ! ROH þ RCHO þ O2 RO2 þ RO2 ! RO þ RO þ O2

2.2

Degradation of Atmospheric Alkenes

In the case of alkenes OH radicals preferably react by addition to the C=C double bond as illustrated for ethene: CH2 ¼ CH2 þ OH ! HO
CH2
CH2 This hydroxyalkyl radical reacts with an oxygen molecule to form a hydroxyalkyl peroxy radical, HO
CH2
CH2 þ O2 ! HO
CH2
CH2 O2 which is converted to a hydroxy alkoxy radical following the reaction with NO. HO
CH2
CH2 O2 þ NO ! HO
CH2
CH2 O þ NO2 The hydroxy alkoxy radical can then either react with O2 via H-abstraction to become a glycolaldehyde or by cleavage to form an aldehyde and a hydroxy alkyl radical, which can react again as described above. Furthermore, larger chained molecules have the possibility to undergo isomerization. The second major pathway for the degradation of alkenes is their reaction by addition of an ozone molecule (O3) to the C=C double bond leading to an instable so called ozonide, which then decomposes to a carbonyl compound and a Criegee intermediate, as shown for ethene: CH2 ¼ CH2 þ O3 ! HCHO þ ½H2 COO

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Chemistry of Volatile Organic Compounds in the Atmosphere

Criegee intermediates can either stabilize by collision with other molecules or decompose further. After stabilization, a possible pathway leads via reaction with water vapor to the formation of organic acids. The reaction of alkenes with the nitrate radical also proceeds through the addition of the radical to the double bond. This reaction leads to formation of carbonyls and nitro-oxy carbonyls. These compounds react further to finally form intermediate and stable products, which then are oxidized as described above.

2.3

Degradation of Atmospheric Aromatic VOCs

The degradation of aromatic VOCs proceeds by two different pathways, the abstraction of a hydrogen atom from one of the alkyl substitute groups or by OH-radical addition to the aromatic ring. In the first case, the stable product is an aromatic aldehyde (e.g., toluene leading to benzaldehyde). However, the OH addition is the predominant pathway for the degradation of aromatic VOCs. Following the addition of an OH radical to the aromatic ring, molecular oxygen is added to build a cyclic hydroxy peroxy radical. In the following the ring structure is opened and epoxy compounds, saturated and unsaturated dicarbonyl radicals and finally methylglyoxal are formed. In the last years a large number of laboratory studies improved our knowledge of the atmospheric oxidation of aromatic hydrocarbons. However, for most of these compounds the detailed reaction pathways in the atmosphere, especially the process of ring-opening, as well as the formation of intermediate and final reaction products are still speculative. Also, their potential to form secondary organic aerosols is far from being understood and an important objective of current research.

2.4

Degradation of Atmospheric OVOCs

Oxygenated volatile organic compounds (OVOCs) are either directly emitted by anthropogenic or biogenic sources. For example, they account for the largest part of organic compounds emitted from biomass burning. As mentioned above, they are also formed during the oxidation of VOCs. Again, the predominant reaction is the abstraction of a hydrogen atom from the carbon chain by an OH radical. In the case C=C double bonds are present, also the addition of O3 is also a possible initial step. The resulting peroxy radicals then react as described above. Those OVOCs that have UV-absorbing groups (e.g., aldehydes, ketones, organic peroxides and organic nitrates) can also be photodissociated. Whereas formaldehyde is degraded primarily by photolysis, higher aldehydes react mainly with OH radicals. As shown for formaldehyde, products of the photolysis provide an additional source of HO2, which can then be a source for OH radicals: HCHO þ hv ! CO þ H2 HCHO þ hv ! HCO þ H HCO þ O2 ! HO2 þ CO H þ O2 þ M ! HO2 þ M

Chemistry of Volatile Organic Compounds in the Atmosphere

18

The initial steps of the aldehyde degradation by OH radicals is shown for the example of acetaldehyde: CH3 CHO þ  OH þ O2 ! CH3 CðOÞO2 The produced peroxy radical is a precursor for the PAN formation, if enough NO2 is available (see above). At high NO levels, however, the peroxy radical of a Cn-aldehyde reacts dominantly with NO, leading to a Cn-1-aldehyde and CO2. For the OH-radical initiated oxidation of ketones the reaction proceeds by H-atom abstraction and subsequent formation of alkoxy radicals. Also the oxidation of alcohols in the atmosphere is mainly initiated by the reaction with OH radicals. The H-atom is abstracted from the C–H bond of the CHOH or CH2OH group. The following reaction of O2 with the hydroxyradical leads to the formation of a ketone for secondary alcohols or of an aldehyde for primary alcohols. Furthermore, OH can abstract H-atoms from other C–H bonds, which leads to reactions analogous to those for alkanes.

3

Research Needs

Almost all human activities (even breathing) lead to the emission of organic compounds into the atmosphere. Additionally, the terrestrial vegetation releases huge amounts of organic compounds into the air. If climate changes, as it is predicted to do, temperatures will increase, convection systems and vegetation patterns will alter significantly, and with it most likely the distribution and the composition of organic compounds in the atmosphere. At the same time, the increasing population with a steady growth of urban areas and an increasing number of megacities demand a basic necessity to assess and control air quality in order to maintain human health. All this requires a still better understanding of the sources, sinks and the chemistry of organic compounds in order to improve models for the prediction of future global change and develop appropriate abatement strategies.

References Atkinson R, Arey J ( 2003) Atmospheric degradation of volatile organic compounds. Chem Rev 103: 4605–4438. Calvert JG et al. (2000) The Mechanisms of the Atmospheric Oxidation of Alkenes. New York: Oxford University Press. Finlayson-Pitts BJ, Pitts JN (2000) Chemistry of the Upper and Lower Atmosphere. San Diego, USA: Academic Press.

Seinfeld JH, Pandis SN (2000) Atmospheric Chemistry and Physics. New York: Wiley. Koppmann R (ed.) (2007) Volatile Organic Compounds in the Atmosphere. Oxford: Blackwell Publishing. Zellner R (ed.) (1999) Global Aspects of Atmospheric Chemistry. Darmstadt, Germany: Steinkopf.

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19 Hydrocarbons in the Pedosphere L. Schwark Institute for Geosciences, Christian-Albrechts-University, Kiel, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 280

2 2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.1.5 2.2 2.2.1 2.2.2

Generation within and Transfer of Hydrocarbons into the Pedosphere . . . . . . . . . 281 Natural Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 281 Hydrocarbons Derived from Vegetation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 281 Hydrocarbons Derived from Soil Microbiota and Macrobiota . . . . . . . . . . . . . . . . . . . . 283 Hydrocarbons Derived from Natural Combustion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285 Short-Term Degradation of Plant Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 285 Abiotic Factors Influencing Stabilization of Lipids in Soils . . . . . . . . . . . . . . . . . . . . . . . 287 Anthropogenic Contaminants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 Petroleum-Derived Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 PAH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 288

3 3.1 3.2

Hydrocarbons as Tracers of Recent Environmental Change . . . . . . . . . . . . . . . . . . . . . 288 Molecular Tracers of Land Use and Paleoclimate Change . . . . . . . . . . . . . . . . . . . . . . . . . 289 Sequestration of Atmospheric CO2 in Soils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289

4

Hydrocarbons in Fossil Paleosoils . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 289

5

Molecular Archaeology of Soil and Related Matrices . . . . . . . . . . . . . . . . . . . . . . . . . . . . 290

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_19, # Springer-Verlag Berlin Heidelberg, 2010

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Hydrocarbons in the Pedosphere

Abstract: Organic matter within the pedosphere derives from natural and increasingly from anthropogenic sources. Vegetation comprises the major source of organic matter input into soils, where this input serves as a substrate for soil microbiota and macrobiota, which may constitute up to one third of the biomass in soils. Although the dominant fraction of soil organic matter is macromolecular, free and bound lipids constitute an important compound class of highly diagnostic value for the reconciliation of biological sources and alteration processes. Hydrocarbons and their functionalized analogs may have long residence times in soils allowing for the reconstruction of recent and fossil environmental conditions and changes thereof. Long-chain acylic compounds and cyclic terpenoids derived from cuticular lipids are employed in the reconstruction of land-use change and paleoclimate evolution. Short-chain components, in particular fatty acids, are indicative of soil microbial community structures and biogeochemical alteration processes. Hydrocarbons frequently occur as contaminants in soils due to spillage of petroleum (products) and ubiquitous emissions from fossil fuel or biomass combustion.

1

Introduction

Hydrocarbons comprise an abundant compound class of high structural diversity (See > Chapter 1, Vol. 1, Part 1) that is found in and exchanged between all compartments of the geobiohydrosphere. The pedosphere is one important compartment within the continuum of the geobiohydrosphere and is defined as ‘‘the envelope of the Earth where soils occur and biogeochemical processes are active.’’ Soil development and formation processes are important factors that will regulate the abundance and composition of hydrocarbons found in the pedosphere. The description of soil formation and the various soil types, however, is beyond the scope of this contribution, and the reader is referred to the multitude of excellent textbooks on this subject. Soils do form where an input of organic matter biosynthesized by primary producers enters the surface layer of the terrestrial environment. Although subaquatic soils may form by episodic flooding, e.g., in river floodplains, deltas, marshlands, or in rice paddies, soils are commonly regarded as terrestrial environments. The input of primary biomass serves as a substrate for highly variable soil macrobiota and microbiota, whose activities are required for soil formation. Primary biomass input predominantly derives from higher plant vegetation, such as trees, shrubs, herbs, and grasses and occurs mainly in the form of litter and wood, and also as rhizodeposition that is estimated to account for up to 30% of vegetational organic matter input into soils. Before the development of higher land plants in the Late Silurian (first primitive land plants) to Early Devonian (first forest ecosystems), i.e., some 450–360 (million years before present [MaBP]), algae, fungi, and lichens had already colonized land surfaces and initiated soil formation (Algeo and Scheckler, 1998). As soon as the vegetation input becomes available as substrate for utilization, further soil biota will develop. Macrofaunal biota, e.g., insects and worms, will hydrolyze and break down vegetation input into a size digestible by microbiota. In Section 2.1, soil hydrocarbons derived from vegetation and microbial communities in soil are described, and their diagenetic transformation is discussed. Since neolithic times, anthropogenic perturbations of natural soil formation and hydrocarbon input into the pedosphere have been occurring. In particular, the combustion of biomass and, since the onset of the industrialization, of fossil fuels or their processing products has led to

Hydrocarbons in the Pedosphere

19

a dominantly anthropogenic fraction of hydrocarbon input into soils. In Section 2.2, a brief summary of anthropogenically derived hydrocarbons in soils is given with a focus on hydrocarbons sensu strictu, i.e., aliphatic and aromatic hydrocarbons. Xenobiotic components (pesticides, plasticisers, pharmaceuticals, personal care products, etc.) are excluded from the discussion, but covered in other chapters of this series. In Section 3, a brief summary is given about how soil hydrocarbons may be applied in the reconstruction of fairly recent, i.e., within the past centuries, environments and environmental changes. This includes soil hydrocarbons as tracers of paleoclimate, paleovegetation, and landuse change. In addition, the role of soil hydrocarbons in CO2 cycling and sequestration is addressed. Fossil soils or paleosols have been described back to the Proterozoic, corresponding to older than 560 MaBP, i.e., they predate the establishment of higher plant communities on land by hundreds of millions of years. During the geological record, major perturbations have occurred leaving marked traces in soil hydrocarbon composition. These have been used to elucidate mechanisms potentially responsible for environmental changes as discussed in > Chapter 3, Vol. 1, Part 1. The last part of this summary briefly addresses the utilization of soil hydrocarbons in the archaeology or archaeometry, in particular, early forms of land management and cultivation.

2

Generation within and Transfer of Hydrocarbons into the Pedosphere

2.1

Natural Compounds

2.1.1

Hydrocarbons Derived from Vegetation

Land plants are complex life forms constituting an enormous variety of chemical components. The major compound classes found in living plants serve for structural integrity, like proteins, celluloses, and lignin, or for energy storage, like starch or proteins. They occur as macromolecular components that only upon degradation by soil biota will release hydrocarbon moieties into the pedosphere. This chapter addresses primarily the occurrence of hydrocarbon lipids in terrestrial plants, their chemotaxonomic and, thus, (pal)ecological value, and their transformation along the pathway from living plant via senescent plant to litter and the humus layer and finally through various soil horizons. Terrestrial plant lipids are predominantly located in the cell membrane, the cuticle layer, intracuticular compartments, epicuticular waxes, and plant oils and resins. Such lipid compounds may occur in various aboveground and belowground plant organs, including flowers, leaves, needles, blades, stems, barks, and roots. These plant compartments constitute the main interface layer between the plant and its surrounding environment, the atmosphere, hydrosphere, and pedosphere. Exchange of fluids has to be maintained permanently between the plant and its environment under conditions highly variable in, e.g., temperature, humidity, electromagnetic radiation, and gas composition, as well as pressure or concentration (Riederer and Muller, 2006; Samuels et al., 2008). Cuticles and cuticular waxes, thus, have to fulfill a variety of tasks and consequently are highly complex and variable in composition. The cuticle is composed of the macromolecular polyesters cutin that forms a structural backbone in which free-extractable intracuticular waxes are embedded (Kolattukudy, 1980; Samuels et al., 2008). The cutin polyester is composed of terminal- and mid-chain hydroxy

281

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fatty acids, mainly in the C16–C18 range and with 9,16- and 10,16-dihydroxyhexadecanoic acid and 9,10,18-trihydroxyoctadecanoic acid being of diagnostic significance. In addition, alkanoic monoacids and diacids, as well as n-alcohols and alkanediols (Ko¨gel-Knabner, 2002; Kolattukudy, 1980, 2001) are important cutin building blocks. The second most important polyester in plants is suberin, which is present in cork cells of tree bark or in epidermal cells of grass, but quantitatively most importantly it covers subterranean root surfaces. Suberin differs from cutin in that it is composed primarily of longer chain, preferentially terminal n-alkanols and alkanoic acids in the C20–C30 range and dicarboxylic acids with C24 often predominating. In contrast to cutin, suberin may contain aromatic moieties. Cutin and suberin are highly recalcitrant and will release their monomeric constituents into soils only over prolonged periods of time, mainly via depolymerization by extracellular fungal cutinases. A substantial proportion of C12–C30 functionalized hydrocarbons in soils may, thus, be attributed to the enzymatic depolymerization of cutin and suberin polyesters (Ko¨gel-Knabner, 2002; Kolattukudy, 2001; Nierop et al., 1998). For specific groups of plants, mainly xerophytic CAM plants like Agave americana, the presence of the non-hydrolyzable highly cross-linked aliphatic biopolymers, cutan and suberin, has been proposed (Nip et al., 1986; Tegelaar et al., 1989, 1995) that upon degradation may also yield hydrocarbons to the soil lipid pool. The intracuticular waxes that typically constitute 20–60% of the cuticle mass (Heredia, 2003) are rapidly released into the pedosphere and form an easily solvent-extractable fraction of the total soil organic matter. Epicuticular waxes cover the lamellar outer cuticle proper and often build up a protective barrier around stomatal antechambers. Although generally similar to intracuticular lipids the extracellular waxes may differ in chemical composition (Gniwotta et al., 2005; Jetter et al., 2006) and in their crystallinity or surface properties (Neinhuis and Barthlott, 1997). Comparable to the macromolecular lipidic polyesters the intra and epicuticular waxes show a high degree of structural variability (Jetter et al., 2006; Samuels et al., 2008) with substantial chemotaxonomical potential. The main structural classes of land plant wax lipids encompass long-chain (C>20) acyclic components like n-alkanes, iso-alkanes, n-aldehydes, n-alcohols, n-carboxylic acids, ketones, and esters, and cyclic components, mainly triterpenoids. The distributional pattern of n-alkanes due to their diagenetic stability has been used frequently for chemotaxonomic separation (e.g., Dove, 1992; Eglinton et al., 1962; Herbin and Robins, 1968; Jansen et al., 2006; Maffei, 1996a, b; Piasentier et al., 2000; Rommerskirchen et al., 2006; Schwark et al., 2002; Vogts et al., 2009; Zhang et al., 2004) though adaption of wax composition due to seasonal variation in temperature, water and nutrient availability, bioproductivity, irradiation, CO2 concentration, grazing, insect attack, or other environmental stress and various physiological factors (Barnes et al., 1996; Cameron et al., 2002; Dodd and Poveda, 2003; Dove et al., 1996; Jenks, 2001; Prasad and Gu¨lz, 1990; Rieley et al., 1993, 1995; Teece et al., 2008; Wiesenberg et al., 2008a, b; Zhang et al., 2004) have to be considered. Iso-alkanes commonly constitute trace components in plant lipids but may be diagnostically enriched in tobacco or potato plants (Barnes et al., 1996; Szafranek and Synak, 2006). Functionalized lipids easily and early undergo structural modification upon plant senescence and litterfall (Nguyen Tu et al., 2001) continued by soil lipid formation. Substantial variation of wax lipid composition among species has been reported and related to surface properties, e.g., glaucousness (Jenks, 2001; Osborne and Stevens, 1996). Chemotaxonomy has been based on

Hydrocarbons in the Pedosphere

19

carboxylic acids (Mongrand et al., 1998, 2001), primary and secondary alcohols (Franich et al., 1979; Gu¨lz, 1994; Jetter et al., 2006), diols (Gu¨lz, 1994; Jetter et al., 2006), esters (Su¨mmchen et al., 1995), or estolides (Gu¨lz, 1994). For a comprehensive review of plant wax lipids see Jetter et al. (2006). Furthermore, acyclic n-alcohols and carboxylic acids have been used for discrimination of C3/C4-plants (Bianchi and Bianchi, 1990; Rommerskirchen et al., 2006; Vogts et al., 2009; Wiesenberg and Schwark, 2006). The ratio of C24/(C22 + C26) carboxylic acids in C4 crop plants is higher than that in C3 plants (Wiesenberg and Schwark, 2006). According to Vogts et al. (2009), the maximum in the distribution pattern for n-alkanols of rainforest plants (n-C30 alkanol) lies between those of C3 savanna plants (n-C28 alkanol) and C4 grasses (n-C32 alkanol). A substantial database exists on the composition of lignin and its oxidative chemical or pyrolytic breakdown products in soils (Rasse et al., 2006). Differentiation of gymnospermal versus angiospermal lignin is achieved via the distribution of syringyl moieties derived from angiosperms and vanillyl or guajacyl groups stemming from gymnosperms. Differentiation of tissue types is achieved by the relative abundance of cinnamyl groups that originate from breakdown of nonwoody lignin (Goni and Eglinton, 1996; Hedges and Mann, 1979).

2.1.2

Hydrocarbons Derived from Soil Microbiota and Macrobiota

The composition of phospholipid fatty acids (PLFAs) has been intensively used to study microbial biomass activity and community structure in soils because PLFAs are only found in viable cells and hence are characteristic biomarkers for living microorganisms (Evershed et al., 2006; Frostega˚rd and Ba˚a˚th, 1996; Zelles et al., 1992; Zelles, 1999; Webster et al., 2006). Structural variability of PLFAs, in principal their methyl-branching position, the insertion of cyclopropane rings and the occurrence, as well as the position of one to three double bonds can be used to establish the notional proportions of fungi, gram-positive bacteria (including actinomycetes), or gram-negative bacteria (Frostega˚rd and Ba˚a˚th, 1996; Zelles, 1999). Phenotypic profiling techniques such as PLFA analysis aim toward the analyses of microbial membrane composition and hence, do not distinguish between microbial species (Singh et al., 2006). The application potential, thus, primarily lies in the determination of the relative proportions of fungi, gram-negative, and gram-positive bacteria in soils. Source-diagnostic PLFA patterns in the literature have been compiled (Amelung et al., 2009; Feng and Simpson, 2009; Frey et al., 2008; Frostega˚rd and Ba˚a˚th, 1996; Kieft et al., 1994; Zhang et al., 2005; Zelles et al., 1992). The utilization of soil or litter carbon pools occurs via specialized groups of microorganisms, typically resulting in a cascade of microbial processes during decomposition. Hereby, fungal growth is generally found to dominate during early stages of plant residue decomposition (Feng et al., 2007), and in the mineralization of coniferous plant litter (Berg et al., 1998). Gram-positive bacteria are abundant in soils with low substrate availability and in subsoils with lower organic carbon content (Griffiths et al., 1999). In contrast, gram-negative bacteria prefer an input of fresh organic material (Griffiths et al., 1999; Kramer and Gleixner, 2006). In a recent review, Joergensen and Wichern (2008) summarize the diagnostic potential of PLFA for discrimination of fungal biomass in soils. Actinomycetes are identified by esterlinked 10-methyl branched saturated fatty acids (McKinley et al., 2005; Williams and Rice, 2007). Saprotrophic and ectomycorrhizal fungi from the phyla Zygomycota, Ascomycota, and Basidiomycota are characterized by the presence of linoleic acid (18:2o6). Oleic acid (18:1o9)

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and g-linolenic acid (18:3o6) have been used as general fungal biomarkers (Baath and Anderson, 2003; Potthoff et al., 2006). However, oleic acid (Ruess et al., 2007) and g-linolenic acid (Arao et al., 2001) have also been employed to discriminate plant and colembolla contributions to soil PLFA. As an indicator for arbuscular mycorrhizal fungi (phylum Glomeromycota), 16:1o5 has been successfully applied (Olsson et al., 1995). It must be taken into consideration that at present no conversion factors are available for total PLFA concentration to soil bacterial carbon or any specific PLFA to gram-negative bacteria, or gram-positive bacteria, such as Firmicutes and Actinobacteria. Different assignations for bacterial PLFA were used, for example, by Gattinger et al. (2002) and Marschner et al. (2003), indicating the urgent need for more basic data on bacteria. A means to improve further the power of PLFA analysis will be by the increasing application of compound-specific d13C fatty acid analysis (Glaser, 2005; Kramer and Gleixner, 2006; Olsson et al., 2005). Although not strictly related to functional traits, the relative abundance of fungi, gramnegative, and gram-positive bacteria is informative of the microbial community composition in soil (Frey et al., 2008; Zhang et al., 2005), and may be applied to indicate microbial community shifts. Such changes in microbial consortia may be induced by changes in a number of different environmental variables including among others temperature, pH, Eh, substrate limitation, or presence of metal toxins (Feng and Simpson, 2009; Feng and Simpson, 2007; Frostega˚rd and Ba˚a˚th, 1996; Zelles et al., 1992). Fungal lipids in soils are well assessed by the presence of ergosterol, a highly specific C28-steroid predominant in fungal cell membranes. Ergosterol is endogenous only in higher fungal phyla, i.e., Basidiomycota, Ascomycota, and the majority of Zygomycota (Klamer and Baath, 2004; Niemenmaa et al., 2008; Weete and Ghandi, 1996, 1999). Over the past years Ergosterol has, thus, been increasingly used as a biomarker for the presence and activity of fungi in soil (Grant and West, 1986; Montgomery et al., 2000; Rousk and Baath, 2007; Yergeau et al., 2007). The widespread presence of archaea in various environmental compartments has only recently been recognized (Schleper et al., 2005). In soils, archaea have been first shown to occur by combined application of lipid analyses coupled to molecular genetics by Gattinger et al. (2002). Their important role in nutrient cycling, in particular during ammonium oxidation, has recently been demonstrated by Leininger et al. (2006). Due to their unique membrane building blocks, crenachaeota can be detected by their specific glyceroldialkylglyceroltetraethers (GDGT) of the crenarchaeol type. A correlation between crenarchaeol lipid concentration and ammonium monooxygenase gene copies in the studied soils indicates that most soil archaea in those samples may be nitrifying archaea (Leininger et al., 2006). Further studies on the role of archaea in terrestrial environments are needed to fully comprehend their biogeochemical effects on soil organic matter. A second class of GDGT with an isoalkane (branched isoalkane tetraethers [BIT]) instead of isoprenoidal dialkyl structure was subsequently discovered (Weijers et al., 2006) that could be linked to ubiquitous soil bacteria. The distribution of these bacterial GDGT in soils is controlled by prevailing pH conditions and soil temperature (Weijers et al., 2007). Increased cyclization of BIT preferentially occurs in soils of lower pH, whereas increased methyl branching of the isoalkane chains corresponds with lower soil temperature (Weijers et al., 2007). Investigating the distribution of GDGT now allows for an improved characterization of environmental variables in terrestrial systems, in particular including soils.

Hydrocarbons in the Pedosphere

2.1.3

19

Hydrocarbons Derived from Natural Combustion

The discrimination of natural and anthropogenic combustion residues embedded in soils often is neither easy to achieve nor practical (Ribes et al., 2003; Tinoco et al., 2006). For simplicity, the compounds derived from combustion of natural biomass will be covered here, and the residues derived from fossil fuel combustion will be covered in Section 2.2. Natural or man-made fires release abundant lipid components into the biosphere, in particular, if burning conditions favor the only partial destruction of biomass (mainly dehydration) over the complete destruction to char or ‘‘black carbon.’’ Smoke may transport lipid residues over long distances, but finally deposition in soils or sediments will occur (Simoneit, 2002). The spectrum of compounds released into the atmosphere mainly depends on the primary biochemicals of the burning vegetation, e.g., grass, shrubs, conifers, angiosperms, and the intensity of combustion. Steam distillation processes and volatilization of low-molecular compounds at low temperatures (400 C) lead to the formation of highly condensed polycyclic aromatic hydrocarbons (PAHs) of limited diagnostic source value (Simoneit, 2002; Wiesenberg et al., 2009). Smoke contains a broad variety of long-chain n-alkanes, alcohols, carboxylic acids or diacids, and wax esters inherited from primary plant wax constituents (Oros and Simoneit, 2001). In addition, monoterpenoids, sequiterpenoids, and diterpenoids from vegetation may be abundant or even dominant smoke constituents (Oros and Simoneit, 2001). Phenols formed by the degradation of lignin and levoglucosan and other sugar derivatives formed by the degradation of cellulose and hemicellulose are typical combustion markers. While combustion-derived waxes and terpenoids are stable compounds that after deposition become embedded into soil organic matter, sugars are easily metabolized and often do not accumulate in soils. Fresh charred materials may, however, still contain significant amounts of burnt cellulose (levogucosan, mannosan, or galactosan) as shown by Otto et al. (2006) at recent wildfire sites in Canada. The presence of n-alkenes, sterenes, and other unsaturated hydrocarbons in soils due to their high reactivity can be taken as an indication of fresh biomass combustion, which in a moderate temperature regime will yield alkenes by dehydration (Simoneit, 2002; Wiesenberg et al., 2009). The PAHs formed from biomass under harsher combustion conditions as in forest fires or upon volcanic activities (Ribes et al., 2003) usually appear as parent PAH (Simoneit, 2002; Tinoco et al., 2006), which is in contrast to long-lasting formation under geological conditions, which will yield preferentially alkyl-substituted PAH (Simoneit, 2002). It is generally assumed that almost the entire proportion of PAHs found in soils does derive from either natural or anthropogenic combustion sources. Only in rare occasions, eolian PAH input from coal-bed erosion may occur. In tropical regimes, it has been postulated that low-molecular weight PAHs like naphthalene and phenanthrene may originate from termites or their associated fungi (Krauss et al., 2005). Perylene is a five-ring PAH for which also a diagenetic formation in sediments as soils has been inferred based on concentration profiles and carbon isotopic composition (Wilcke et al., 2002).

2.1.4

Short-Term Degradation of Plant Lipids

Degradation of plant-derived lipids may occur as early as upon leaf senescence (Nguyen Tu et al., 2001) but latest during litter formation. Functionalized plant lipids such as

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alcohols, carboxylic acids, and wax esters embedded in litter layer and soils pass through a diagenetic continuum of aerobic and anaerobic, enzymatically and non-enzymatically transformation reactions, mediated by soil macrofauna and/or microorganism. Physicochemical boundary conditions and processes, e.g., aggregate protection, reaction with polyvalent cations, pH, clay content, and strong partitioning between water phase and mineral surfaces serve to enhance soil lipid preservation in soils. Upon waterlogging and under low pH conditions, e.g., during podsolization, microbial activity in soils is retarded (van Bergen et al., 1998) and free lipids may contribute up to 20% of the total soil organic matter (Bardy et al., 2009; Naafs et al., 2004a, b; Nierop et al., 2005; Que´nea´ et al., 2004, 2006; van Bergen et al., 1998). Aliphatic lipids are known to biodegrade in soils and surface sediments mainly via oxidation first to n-alkanols and then ultimately to their corresponding n-alkanoic acids (Amble´s et al., 1994a, b; Dinel et al., 1990; Otto and Simpson, 2005). Long-chain carboxylic acids can be easily metabolized by numerous aerobic bacteria via beta oxidation resulting in the formation of shorter n-alkanoic acids (Amble´s et al., 1994a; Dinel et al., 1990), which will be utilized as substrate by soil microorganism. Ketones may enter the soil lipid pool directly as constituents of plant waxes or as methylketones/ethylketones resulting from n-alkane or n-alcohol degradation producers, whereby climatic and soil microbial community structures may affect the distribution of methylketones versus ethylketones (Ambles et al., 1993; Bai et al., 2006; Jansen and Nierop, 2009). With increasing degree of degradation, a shortening in the chain length of acyclic aliphatic lipids is observed pointing toward stepwise decarboxylation of the primary plant wax compounds (Bardy et al., 2009; Dinel et al., 1990; Otto and Simpson, 2005). Alternatively, n-alcohols and carboxylic acids may undergo random polymerization and, thus, contribute to humification and the formation of stabilized macromolecular soil organic matter (Ko¨gel-Knabner, 2002; Marschner et al., 2008). Information about the fate of cyclic lipids such as steroids and pentacyclic terpenoids in soils is limited (Otto and Simpson, 2005; van Bergen et al., 1997) though it has been shown that the suite of aliphatic lipids (n-alkanols, n-alkanoic acids, n-alkanes, and wax esters) appears to be preferentially degraded compared to the cyclic steroids and terpenoid lipids (Otto and Simpson, 2005). Hopanoids, derived from soil heterotrophic bacteria or cyanobacteria and other pentacyclic triterpenoids of the lupane, ursane, olanane type derived from higher angiosperm plants significantly contribute to the soil lipid pool (Bull et al., 2000; Jaffe´ et al., 1996; Naafs et al., 2004a, b; Ries-Kautt and Albrecht, 1989; Shunthirasingham and Simpson, 2006; van Bergen et al., 1997). Bacteriohopanepolyols have only very recently been analyzed in soils due to progress in LC-MS analytical techniques (Cooke et al., 2008; Xu et al., 2009). In grassland soils adenosylhopane was identified as specific marker for soil bacteria. The concentrations determined for highly functionalized bacteriohopanepolyols outweigh those of the free-extractable defunctionalized hopanoids by a factor of 100 and more, thus, underlining the importance of soil microbial biomass. Most pentacyclic triterpenoids found in soil are derived from angiosperm leaves or roots (e.g., Bardy et al., 2009; Naafs et al., 2004a, b; Nierop et al., 2005; van Bergen et al., 1997). In soils, free-extractable sterenes and pentacylic triterpenes are usually easily degraded or incorporated into the bound lipid fraction leading to a rapid concentration decrease with depth (van Bergen et al., 1997), except if an important input by roots occurs, as described for podsolization (Naafs et al., 2004a). In most soils, functionalized triterpenoids are stabilized by progressive aromatization (Bardy et al., 2009; Otto and Simpson, 2005; Ries-Kautt and

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Albrecht, 1989). In forest soils the bicyclic (cadalenes) and tricyclic (abietanes, pimaranes, and isopimaranes) aromatic hydrocarbons of sesquiterpenoid and diterpenoid origin derived from conifers can be abundant (Jaffe´ et al., 1996; Otto and Simpson, 2005).

2.1.5

Abiotic Factors Influencing Stabilization of Lipids in Soils

A recent review by Von Lu¨tzow et al. (2006) summarizes evidence for abiotic mechanisms operating in soils that will contribute to preserve organic matter. They conclude that primary recalcitrance of organic matter, which includes plant litter, rhizodeposits, microbial products, humic polymers, and charred residues, is of less importance for preservation. Of critical importance for preservation, they regard the spatial inaccessibility of a large variety of soil organics against decomposer organisms due to occlusion, intercalation, hydrophobicity, and encapsulation and further, the stabilization by interaction with mineral surfaces (Fe-, Al-, Mnoxides, and phyllosilicates) and metal ions. With respect to the protection of lipid hydrocarbons in soils the dominant mechanism is thought to be occlusion in soil macroaggregates >250 mm, which is further promoted by the hydrophobicity of long-chain aliphatic moieties in n-alkanols, carboxylic acids, and n-alkanes. The role of metal lipid interactions for preservation of the latter is assumed to be of minor importance only.

2.2

Anthropogenic Contaminants

Hydrocarbons have been released into the environment in vast amounts by anthropogenic activities, in particular since the industrial revolution and the large-scale exploitation of fossil fuels (coal, oil, and gas). Reviews of the type of components released and their current inventories in various environmental compartments are available, with several studies focusing on hydrocarbons in soils.

2.2.1

Petroleum-Derived Compounds

Petroleum constitutes a multicomponent and highly variable mixture of hydrocarbons (Peters et al., 2005), often dominated by n-alkanes, branched alkanes, and cycloalkanes covering a broad range of molecular sizes from approximately 7–100 carbon atoms (for structures, see > Chapter 1, Vol. 1, Part 1). Aromatic and naphtheno-aromatic hydrocarbons, as well as heterocompound (O, N, S) bearing components are present in usually smaller amounts (Peters et al., 2005). Release of petroleum or petroleum products is the main mode of hydrocarbon enrichment in the pedosphere. Thermal maturation of petroleum generates an even distribution in the chain length of its n-alkanes with a carbon preference index (CPI, see Peters et al., 2005 for further details) around 1.0. In contrast, input of fresh n-alkanes derived from vegetation into soils is strongly dominated by long-chain odd-numbered homologues like nonacosane or hentriacontane. The relative concentration of the n-alkane homologues in agricultural soils may depend on the crops grown in the surrounding (Conte, 2003; Wiesenberg et al., 2004) or in forests depend on the prevalent tree species.

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Microbial degradation of petroleum alkanes leads to the formation of an unresolved complex mixture (UCM) that is found in sites with former petroleum contamination (Peters et al., 2005). A variety of diagnostic tools to decipher the origin of even strongly degraded petroleum in natural matrices is available (Faure et al., 2007; Peters et al., 2005; Wang et al., 2006). In particular, two-dimensional gas chromatography is capable of resolving several thousands of individual hydrocarbons in UCM samples (Frysinger et al., 2003), thus, enabling source typing. The toxic potential of petroleum hydrocarbons for soil biota due to limited bioavailability is commonly considered low (de Jonge et al., 1997) though elevated concentrations of PAH in spilled petroleum may be critical.

2.2.2

PAH

A summary of general PAH occurrence in soils and their respective pathways of introduction into soils are given by Wilcke (2000). Detailed inventories of PAH in soils have been presented on a global scale (Wilcke, 2007) and for various regions, e.g., China (Cai et al., 2008; Ping et al., 2007), the UK (Wild and Jones, 1995), France (Motelay-Massei et al., 2004), Spain (Nadal et al., 2004), or Russia (Wilcke et al., 2006). Published data on PAHs patterns in topsoils from 30 regions all over the world (Cai et al., 2008; Wilcke, 2007) identify two distinct PAH patterns, one representing a general background, the other indicating anthropogenic pollution, in particular via incomplete combustion of fossil fuels or recent biomass. The background signature is dominated by two- and three-ring PAH and by perylene, which although being a high-molecular weight PAH is mainly derived from diagenetic conversion of natural precursors (Wilcke, 2000). Composition of PAH mixtures in soils can be explained as simple mixing of these two major source groups. Other factors influencing the composition of PAH mixtures such as transboundary atmospheric redistribution and PAH point sources including different combustion materials and processes may affect soil PAH patterns (Wilcke, 2007). Despite the broad distribution, the multiple sources, and the detrimental health effects of PAH, only limited knowledge is available for remote areas including, e.g., the polar and subpolar regions, tropical Africa and Southeast Asia, and southern hemispherical temperate zones. Reconstructions of PAH accumulation histories in soils have been achieved by using archived soil samples (Jones et al., 1989), thus, eliminating the problem of poor time resolution in natural soil sequencers. In agreement with studies in aquatic sediments, a decline in the overall PAH burden following a maximum in the 1960s due to intensive coal combustion with minimal filtering efforts can be recognized.

3

Hydrocarbons as Tracers of Recent Environmental Change

Stratigraphic or temporal resolution of soil sequences is critically low when compared to sediments accumulating in lakes or marine sedimentary basins. Therefore, due to much better time control, trends in environmental evolution are most often followed by analyses of sediments rather than soils. However, in a few exceptional cases, the temporal resolution in soil profiles maybe sufficient for temporal interpretation. Soil sequences deposited over

Hydrocarbons in the Pedosphere

19

millennia or even millions of years are found in loess deposits, e.g., in USA or in China, and can be traced for paleovegetation or paleoclimate development (see section 4).

3.1

Molecular Tracers of Land Use and Paleoclimate Change

Paleovegetation change in alpine environments, in particular altitudinal deviations from current position of the tree line can be investigated by analyses of soil lipids. As demonstrated by Jansen et al. (2006, 2008) in the high Andes, changes in vegetation patterns can be deducted from n-alkane, n-alcohol, and methyl ketone profiles in soils. Based on the soil hydrocarbon composition, the authors were able to demonstrate an increase in the upper tree line over the past 3,500 years (Nierop et al., 2007; Jansen and Nierop, 2009) which not only was unexpected, but also impossible to prove using classical palynological techniques. In a similar manner, Liu et al. (2005) and Gu et al. (2007) deciphered paleoclimatic control on soil lipid distributions in oxisols recovered from river terraces in southern China. They found pronounced variations in the chain length distribution of n-alkanes and in the isoprenoid patterns for glacial and interglacial stages that could be correlated with the marine isotope record.

3.2

Sequestration of Atmospheric CO2 in Soils

A large number of studies have addressed the question whether soils may serve as sinks or sources for atmospheric CO2. In this respect, the determination of carbon turnover times for several compound classes encountered in soils has proven to be informative (Amelung et al., 2009; Marschner et al., 2008). While it was previously assumed that lipid hydrocarbons have higher turnover rates or shorter residence times in soil as compared to polyphenolic lignin constituents, it was recently shown that the turnover times for both compound classes are in the range of only several decades (Amelung et al., 2009; Rasse et al., 2006; Wiesenberg et al., 2004). The residence times of carboxylic acids have been shown to be slightly shorter than those of n-alkanes (Wiesenberg et al., 2004) due to higher microbial breakdown and subsequent incorporation into humic soil matter. It may thus be assumed that CO2 sequestration may only be effective, if soil lipids are translocated into deeper soil horizons where lower mineralization rates prevail (Marschner et al., 2008). Based on current evidence derived from FACE experiments, Wiesenberg et al. (2008a, b) concluded that differential response of investigated plant species to expectedly elevated CO2 concentrations leads to fairly stagnant turnover times for plant and soil lipids, whereby n-alkanes, n-alcohols, and n-carboxylic acids were less effectively degraded under enhanced CO2 conditions.

4

Hydrocarbons in Fossil Paleosoils

Paleosoils, although shown to occur throughout the entire Phanerozoic geological record and, thus, even predating the colonization of land surfaces by higher plants (Algeo and Scheckler, 1998), reveal little potential for preservation of extractable lipid hydrocarbons. Most studies on paleosoils, thus, focus on the bulk isotopic composition of pedogenic carbonates and bulk organic matter. In contrast to marine and limnic sediments, soils have been exploited as archives of environmental conditions to a much lesser degree. Investigations of soil-embedded

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lipids are restricted to the Cenozoic and preferentially address loess sequences, in particular in China, where such deposits are widespread and occur across large climatic gradients. Soil hydrocarbons structural and isotopic analyses in the Chinese Loess Plateau show that C4/C3 plant ratios determined from n-alkane 13C isotope compositions were lower during glacial stages, but increased during interglacials when the summer monsoon intensified (Freeman and Colarusso, 2001; Zhang et al., 2003; Liu et al., 2005). Soil lipids revealed that the C4/C3 plant ratios are more strongly affected by East Asian summer monsoon intensity other than by atmospheric pCO2 levels in the Chinese Loess Plateau. The expansion of C4 grasses during the Miocene has been demonstrated using molecular and isotopic evidence by Freeman and Colarusso (2001) in parallel to soils and marine sediments that received substantial terrigenous organic matter input.

5

Molecular Archaeology of Soil and Related Matrices

Soils serve as important archives for archaeological artifacts and by their soil lipid composition may reveal information on (pre)historic human utilization (Simpson et al., 1999). In a recent review Evershed (2008) summarizes the application of organic residue analyses in archaeology. A variety of hydrocarbons found in soil may serve as indications of anthropogenic input of manure and other organic fertilizers (Bull et al., 2001, 2003). These biological markers include coprostanol and a series of bile acids that have been detected in soil and historic wastewater canals. Based on the relative proportion of individual bile acids an origin from porcine or ruminant sources can be differentiated (Bull et al., 2003). In addition, human influence on soil development can be traced by investigation of soil hydrocarbon (Bull et al., 1999) and an identification of the crop types grown by historic farm communities maybe determined (Jacob et al., 2008).

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20 Organic Matter in the Hydrosphere J. Schwarzbauer Lehrstuhl fu¨r Geologie, Geochemie und Lagersta¨tten des Erdo¨ls und der Kohle, RWTH Aachen University, Aachen, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298

2 Terrestrial Surface Water Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298 2.1 Transfer and Transport Processes Affecting the Residence of Organic Matter in Rivers and Lakes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 298 2.2 Natural Organic Substances . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 300 2.3 Anthropogenic Organic Contamination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 302 3 Ground Water . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 306 3.1 Influence of Redox-Conditions on Organic Matter Quality in Ground Water . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 306 3.2 Natural Organic Substances . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 307 3.3 Anthropogenic Organic Contamination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 307 4 4.1 4.2 4.3

Marine Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 309 Occurrence and Fate of Organic Compounds in the Marine Environment . . . . . . . . 309 Natural Organic Substances . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310 Anthropogenic Organic Contamination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 310

5

Research Needs and Outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 312

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_20, # Springer-Verlag Berlin Heidelberg, 2010

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Organic Matter in the Hydrosphere

Abstract: Organic matter appears in all compartments of the hydrosphere of both natural and more and more anthropogenic origin. Aquatic organic matter exhibits a high structural diversity and corresponding physico-chemcial properties. Marine and terrestrial surface water bodies including their corresponding sediments as well as groundwater are affected by natural compounds of autochthonous origin from aquatic species, in particular from phyto- and zooplankton, and autochthonous material from terrestrial biota. Anthropogenic pollutants are released to aquatic ecosystems mainly as the result of municipal, industrial, agricultural emissions as well as shipping activities. Dominant factors controlling the fate and distribution of organic compounds in the hydrosphere are partition processes between water phase and particulate matter separating more or less hydrophilic from lipophilic substances. With respect to hydrocarbon chemistry an enhanced geoaccumulation of non functionalized aliphatic and aromatic hydrocarbons in the benthic systems has to be stated.

1

Introduction

Organic matter plays an important role in aquatic ecosystems comprising surface water systems as well as groundwater reservoirs. The high diversity of organic compounds with respect to their structure and the related physico-chemical properties cause a widespread and highly diverse occurrence of a multitude of organic compounds in aquatic systems. A clearly arranged and systematic description of the occurrence of organic substances in the hydrosphere is impedet by the variety of different aspects provoking the transport, transfer and transformation of organic matter. The superimposition of vertical and horizontal fluxes within most aqueous systems, different oxygen availability under anaerobic and aerobic ambiences, the different conditions in marine and terrestrial aquatic systems, transfer processes between the more lipophilic particulate matter and the polar water phase depending on the individual chemical nature of the substances, the molecular size of contaminants, their different emission sources (anthropogenic vs. natural as well as autochthone vs. allochthone), and the huge variety of abiotic and biotic transformation in the different compartments of the hydrosphere effect a singular fate of each individual organic compound in each individual aquatic compartment. However, this chapter represents the attempt to give a brief overview on principals regarding the occurrence and the molecular characterization of organic matter in the aquatic environment of both natural and anthropogenic origin. Since microbial assisted or initiated transformation processes, affecting dominantly the molecular composition and therefore the compound spectra in the hydrosphere, are subject of many following, very detailed chapters, these aspects has been neglected here.

2

Terrestrial Surface Water Systems

2.1

Transfer and Transport Processes Affecting the Residence of Organic Matter in Rivers and Lakes

The most important process affecting the fate of organic compounds in the aquatic environment are the principal partition processes between the polar water phase and the more lipophilic particulate matter. These processes, dominantly adsorption and desorption, depend mainly on the polarity of the compounds as the result of their chemical structure

Organic Matter in the Hydrosphere

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and the resulting physico-chemical properties (See > Chapter 1, Vol. 1, Part 1) and, consequently, most of the organic substances accumulate either in the water phase or in the particulate matter. A quantitative estimation of the environmental behavior of organic compounds is given by the KOW-values describing the partition behavior under steady state conditions between water and octanol as representative for the lipophilicity of particulate matter. This partition determines the principal transport processes and, consequently, the distribution of the organic contaminants. The separation of dissolved organic matter (DOM) from particulate organic matter (POM) is defined operationally by a filtration pore size of 0.45 mm. Partially, the dissolved fraction is subdivided by introduction of a third phase (> Fig. 1), the so-called colloidal organic matter (COM), which is also analytically defined e.g., with an upper particle size of 0.45 mm and a lowest compound mass of 1kDa (Guo et al., 2003; Kerner et al., 2003). Anymore, a further sub compartment, the so-called biofilms, consisting dominantly of microbes and extracellular organic matter has been taken into account for interpreting the environmental behavior of organic substances (Headley et al., 1998). In lakes transport processes are not as complex as in riverine systems. The distribution of dissolved organic compounds is controlled by diffusive processes superimposed by an inlet and outlet flow. Particle associated transport is dominated by aggregation and horizontal sedimentation (e.g., Berdie et al., 1995). On the contrary river systems are characterized by a much higher dynamic of the flow regime affecting also the mobility and appearance of organic compounds. The partial very high vertical water flow determines the transport of dissolved organic matter as well as the distribution of particle associated organic substances. Their vertical transport in rivers is linked with the transportation of the suspended particulate matter (SPM), where the corresponding organic load is referred to particulate organic carbon (POC). Additionally, saltation and reptation of heavier particles near the ground level have to be considered for matter transport in flow direction of riverine systems. Since the polarity of non-functionalized hydrocarbons are generally low, these compounds appear dominantly particle associated and their horizontal and vertical transport is linked with the mobility of particulate matter. On the contrary, highly functionalized compounds exhibit enhanced dipole moments inducing an elevated water solubility and, consequently, an environmental behavior closely related to the water transport. They contribute to the dissolved

. Figure 1 Size distribution of organic carbon and nitrogen in the Chena river water; PON/POC = particulate organic carbon/nitrogen; CON/COC = colloidal organic carbon/nitrogen, DON/DOC= dissolved organic carbon/nitrogen (adapted from Guo et al., 2003).

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organic carbon (DOC) or dissolved organic matter (DOM). However, the partition between solid and aqueous phase is not strict but a dynamic exchange process as described in many studies e.g., for aminoacids or carbohydrates (Hedges et al., 1994). Interestingly, this partition causes also a further important environmental aspect regarding the fate of organic substances. The water phase including the suspended particulate matter represents an aerobic environment, whereas the sedimentary matter is dominantly more anaerobic. Hence, it results in quite different transformation or degradation pathways in the distinct compartments for most of the organic compounds as the result of dissimilar microbial communities. Noteworthy, a comprehensive review on the environmental behavior of anthropogenic contaminants in the sedimentary matter of freshwater systems has been published recently (Warren et al., 2003).

2.2

Natural Organic Substances

Natural organic matter (NOM) in surface water is composed of autochthonous material as the result of biological activity within the aquatic system (e.g., fresh water algae lipids) and of allochthonous substances derived from the surrounding biosphere (e.g., higher plant derived lipids). For many compounds a clear attribution to these two pools of riverine and lacustrine organic matter can be effected, although also various substances are emitted to surface water systems by both aquatic as well as terrestrial biota. For instance Countway et al. (2007) used structural differences of certain sterols to differentiate plankton derived, autochthones contributions (represented by 24-norcholesta-5,22-dien-3b-ol (brassicasterol), cholesta5,22-dien-3b-ol, cholest-5-en-3b-ol (cholesterol), 24-methylcholesta-5,22-dien-3b-ol, 24-ethylcholesta-5,22-dien-3b-ol) from allochthonous, higher land plant derived material (indicated by 24-methylcholest-5-en-3b-ol (camposterol), 24-ethylcholesta-5,22-dien-3b-ol (stigmasterol) and 24-ethylcholest-5-en-3b-ol) (see > Fig. 2). An important fraction of natural organic matter is the so-called refractory material which occurs with average organic carbon content of 0.5–100 mg C/L (Frimmel, 1998). In contrast to the labile or metabolizable fraction, that is subject to a more or less rapid transformation and degradation, the refractory matter exhibits a prolonged residence time in lakes and rivers and, consequently, contributes in particular to the terrigenous matter entering the marine environment (see section 3.1). Based on chemical analysis the relative global proportion of the labile fraction in the major river systems has been estimated to be around 35% (Ittekkot, 1988). As mentioned before, more functionalized substances appear dominantly in the polar water phase. Non-altered biochemicals exhibit widely functional groups and, consequently, elevated water solubility. Hence, with respect to NOM the water phase exhibit huge amounts of biological molecules derived from aquatic organism or their excretion as well as intact biomolecules from terrestrial species. Examples include macromolecular compounds like poly saccharides or fulvic acids (Repeta et al., 2002) as well as low molecular weight substances as short chain carboxylic acids or amino acids. In contrast the organic matter accumulating in the sedimentary system constitutes by more lipophilic biochemicals (e.g., sterols, fatty acids and alcohols, long chain n-aldehydes, functionalized terpenoids, cyclic di- and triterpenes, phytol, squalene) superimposed by huge amounts of defunctionalized degradation products of biogenic precursors. Well known examples of the latter group of compounds are loliolide and actinidiolide, ionenes, different isomers of phytenes and the saturated phytane, steranes

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. Figure 2 Molecular structures of certain sterols differentiating autochthones contributions (upper part) from allochthonous material (lower part).

and unsaturated derivatives, phaeophorbide and related porphyrines as well as 4,8,12,16tetramethylheptadecan-4-olide. They correspond to the biogenic precursors chlorophyll, carotenoids, sterols and tocopherols (see also Cranwell, 1981; Cranwell et al., 1987; Hedges et al., 1994; Ittekkot, 1988; Prahl and Pinto, 1987; Riley et al., 1991; Schwarzbauer et al., 2000). Similar to the water phase the major proportion of the sedimentary natural organic matter in rivers belongs to the humic substances, that are proposed to be generated by abiotically mediated ‘geopolymerisation’ reactions. The resulting structurally complex macromolecules (See also > Chapter 1, Vol. 1, Part 1) represent an organic pool, that is not only objective of extensive structure analysis or structural discussions (e.g., Esteves and Duarte, 2000; Kumke et al., 1999; Schulten and Leineweber, 1996; Sutton and Sposito, 2005) but appears to be also an important reagent for the interaction with natural occurring or anthropogenic low molecular weight substances and metal ions (e.g., Klaus et al., 1998; Zwiener et al., 1999; Northcott and Jones 2000). An overall calculation of the individual groups of main DOC constituents has been performed e.g., for the White Clay Creek (Pennsylvania, USA). The composition was pointed out to be 75% humic substances, 13% polysaccharides, 2% amino acids (dominantly as peptides) and 18% compounds with a molecular weight less than 100 KDa (Volk et al., 1997). However, the composition of biogenic organic material in surface water systems including its sediments is subject to temporal variations and dynamic partition effects. For example, Hedges et al. (1994) demonstrated for the Amazona river the usefulness of studying the partition of amino acids and carbohydrates between the liquid and the solid phase in order to differentiate biogenic sources and to determine degradation processes (Hedges et al., 1994). Studies on organic matter transported by suspended particulate mater (SPM) of the Godavari

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river or the York river exemplified the annual fluctuations as well as varying POM sources (Countway et al., 2007; Gupta et al., 1997). As additional remark it has to be stated, that the terms describing natural organic matter are not clearly defined. A recent review summarized the aspects on the nomenclature of ‘‘natural organic matter’’ (Filella, 2009).

2.3

Anthropogenic Organic Contamination

Major parts of surface water systems are influenced by human activities resulting inter alia in contamination by organic pollutants. The spectra of contaminants reflects the broad usage and application of synthetic chemicals in the anthroposphere and, therefore, varies depending on numerous aspects related to the catchment area like population density, level of industrialization, extension of agriculture and effectiveness of waste water treatment. Most important emission sources in highly industrialized and densely populated regions are municipal waste water and industrial effluents. As a result of insufficient waste water treatment pharmaceuticals and bactericides (e.g., carbamazepin, clofibric acid, triclosan), personal care products comprising fragrances, repellents and UV-protectors (N,N-diethyltoluamide DEET, galaxolide or tonalide, 4-methoxycinnamic acid 2-ethylhexyl ester), plasticizers (e.g., phthalates, N-butylbenzenesulfonamide NBBS, 2,4,4-trimethylpentane-1,3dioldi-iso-butyrate TPDB), flame retardants (e.g., tris(chloroethyl)phosphate TCEP), or detergent related products (nonylphenolpolyethoxylates, ethylene diamino tetraacetic acid EDTA) have been detected in river and lake water (see > Fig. 3) and partially also in the corresponding sediments (Balmer et al., 2005; Dsikowitzky et al., 2004; Schwarzbauer and Heim, 2005). The knowledge on indicative substances reflecting the contamination by industrial point source emissions is much more restricted due to the high chemical diversity of the individual effluents. Few information on typical industrial contaminants discharged to the surface water systems have been described e.g., for the leather, paper, chemical, pharmaceutical, rubber, dye and petrochemical industry (e.g., Bilgi and Demir, 2005; Brigden, 2004; Castillo et al., 1999; Czaplicka, 2003; Lopez-Grimau et al., 2006; Pinheiro et al., 2004; Rao et al., 1994; Reemtsma et al., 1995). The corresponding substances belonged amongst others to the substance classes of benzothiazoles, nitro compounds, chlorinated benzufuranones, volatile organic compounds VOCs, substituted anilines and maines, alkyl phosphates and chlorinated arenes. Although semi polar water pollutants also appear in sedimentary systems (e.g., Kronimus et al., 2004), riverine and lacustrine sediments are contaminated dominantly by less functionalized compounds. Typical sedimentary pollutants belong to the group of halogenated hydrocarbons of both aliphatic as well as aromatic constitution. Examples of halogenated aromatics include chlorinated benzenes and naphthalenes, polychlorinated biphenyles (PCB), polychlorinated dibenzo-p-dioxines and dibenzofuranes (PCDD/PCDF), and polybrominated diphenylethers. Most of these congeneric mixtures appear as the result of technical or commercial application such as technical additives, flame retardants or lubricants. Further on, they are partially also generated as by-products in industrial synthesis and are, consequently, discharged via industrial emissions as well. Many of these compounds are of high environmental relevance due to their ecotoxicological and toxcicological properties combined with a high stability under natural conditions leading to potential geo- and bioaccumulation. Hence, numerous of them are classified as priority pollutants.

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. Figure 3 Load profiles of two contaminants of the Lippe river (Germany) emitted by non-point sources (carbamazepine) and a point source (triphenylphosphine oxide) (adapted from Dsikowitzky et al., 2004).

Aliphatic substances with a higher degree of halogenation have to be regarded also as sedimentary pollutants. Examples include hexachlorobutadiene of potential industrial origin as well as polychlorinated long chain n-alkanes the so-called chlorinated paraffins widely used as technical additives. In a less specific manner unfunctionalized aliphatic hydrocarbons as well as aromatic hydrocarbons can contribute to the sedimentary pollution. Natural aliphatic compounds (like several n-alkanes, phytenes etc.) can be superimposed by thermally generated petrogenic aliphatics which are characterized, e.g., by a different distribution pattern of the n-alkane homologues. For pollution source apportionment also petroleum specific tricyclic aliphatics, the hopanes, have been used (see > Fig. 4) by differentiating their thermodynamical stable isomers from the biogenic ones (e.g., Faure et al., 2007; Yunker and Macdonald, 2003). Sedimentary aromatic compounds originate dominantly as the result of petrogenic contaminations but also from pyrogenic emissions (Srogi, 2007; Stout et al., 2001). In particular polycyclic aromatic hydrocarbons (PAHs) and their alkylated derivatives are common

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. Figure 4 PAH cross plots for source indicative ratios characterizing more remote locations (right side) and urbanized regions (left side) (adapted from Yunker et al., 2002).

constituents of oil and petroleum related products, but are also synthesis products during incomplete combustion of organic material e.g., as the result of vehicular traffic. Based on these different sources petrogenic compounds are characterized generally as primary contaminants, whereas pyrogenic hydrocarbons are entering the aquatic environment indirectly as the result of deposition of airborne particles or soil erosion. Hence, the latter ones referred to secondary contaminants. In order to discriminate both emission sources and the related pollution pathways several indicative ratios using source specific isomers have been introduced and applied to environmental studies on rivers and lakes as well as on marine systems (Barra et al., 200X; Grigoriadou 200X). Common ratios are comparing individual isomers (e.g., anthracene – phenanthrene; fluroanthene – pyren, chrysene – benz(a)anthracene or 1,7– 2,6-dimethylphenanthrene) or the relationship between parent PAHs and alkylated homologues (phenanthrene or fluoranthene and pyrene contrasted to their methylated derivatives) (e.g., Yunker et al., 2002). As a new aspect related to the contamination by fossil fuels, the contribution of coaly material and it ingredients to the riverine environment is discussed currently (e.g.,

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Curran et al., 2000; Yang et al., 2008). Also for this material its environmental impact has to be suggested to follow by specific PAHs (Stout and Emsbo-Mattingly, 2008). Intensive agricultural activities in rural regions lead to the discharge of agrochemicals like herbicides, insecticides or fertilizers into rivers and lakes (Schwarzbauer et al., 2001; Venkatesan et al., 1999; Zhang et al., 1999). These effluents are to be characterized as diffuse emissions entering the aquatic environment either by soil surface runoff or by the interaction of the surface waters with corresponding agriculturally contaminated groundwater. In particular the environmental occurrence and fate of pesticides in the hydrosphere has been given major attention. With respect to the type of pollutant these contaminations can be roughly divided into generations of pesticide classes, on the one hand the older generation, which includes more persistent compounds (e.g., chlorinated pesticides such as DDT, g-HCH or lindane, hexachlorobenzene HCB etc.). These compounds are characterized by an elevated tendency to geo- and bioaccumulation, which led to a more or less global ban of these substances. On the other hand a new generation of more modern pesticides are characterized by higher microbial degradation rates, lower lipophilicity and less toxicity (for comprehensive information on the toxicity of pesticides to aquatic organism see DeLorenzo et al., 2001). Representatives of such pesticides are based on molecular moieties comprising carbamates and thiocarbamates (e.g., carbendazim, EPTC), phosphates (e.g., malathion, dichlorvos), sulfonylureas (e.g., cinosulforon), triazines (e.g., simazine, atrazine) and further nitrogen, sulfur and phosphorous containing moieties. Noteworthy, these pesticides are objectives of numerous investigations on abioticially transformation processes in surface water areas e.g., by photooxidation or hydrolysis (e.g., Abu-Qare and Duncan, 2002; Lartiges and Garigues, 1995). Comprehensive studies on the environmental appearance and distribution of pesticides in rivers and lakes have been performed world wide on many river systems (for an overview see Schwarzbauer 2005). Beside all typical pollutants described so far, also specific but not necessarily toxic or ecotoxic compounds have been analyzed to differentiate emission sources and to point out the spatial and time-related anthropogenic impact on the aquatic environment. This approach using the so-called ‘anthropogenic marker’ has been applied to riverine as well as estuarine systems and has reflected the anthropogenic burden by fecal steroids (e.g., coprostanol – indicating fecal discharge), detergents and its by-products (e.g., linear alkylbenzenes and its sulfonated derivatives LAB and LAS – reflecting municipal sewage effluents) or rubber additives (e.g., 2- morpholinylthiazol – related to urban surface run-off). A comprehensive overview on the anthropogenic marker approach has been published by Takada and Eganhouse (1998). Such marker compounds, but also other indicative substances or pollutants, have been used to describe not only the spatial distribution and lateral dynamics (temporary deposition and subsequent erosion) of contaminated particulate matter, but also to obtain an retrospective insight into the long term storage of particle associated pollution. These was performed on accumulated sediment deposits as received by undisturbed aquatic sedimentation in estuaries as the final sedimentation area of riverine particulate matter as well as in lacustrine systems. Investigations on the terrestrial sedimentation of fluviatile matter on flood plains and wetlands has been performed to a minor extent. However, all these deposits can act as ecological archives (see > Fig. 5), since radiological dating of the sediment layers in combination with quantitative chemical analyses reveals a detailed record of the riverine and lacustrine pollution histories for preserved particle bound contaminants (e.g., Fox et al., 2001; Gevao et al., 2000; Heim et al., 2006).

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. Figure 5 Vertical distribution of environmental contaminants determined in a dated sediment core of the Rhine River (Germany), (adapted from Heim et al., 2006).

3

Ground Water

3.1

Influence of Redox-Conditions on Organic Matter Quality in Ground Water

Organic compounds introduced to the terrestrial underground can undergo various types of transport or modification/degradation processes. In groundwater systems a vertical flux in the water unsaturated zone as well as a horizontal flux in the water saturated zone has to be stated and, consequently, an associated transport of dissolved and particle bound substances can be observed. Concurrently, corresponding aerobic and anaerobic zones have to be differentiated with respect to the microbial assisted degradation processes of organic compounds. Since the unsaturated zone belongs more to the pedosphere the focus of this chapter lies on the saturated aquifers. However, the composition of organic matter in aquifers is dominantly controlled by soil derived material, which is valid for natural as well as anthropogenic contaminations. Further on, since water flow rates in aquifers are typically low as compared to rivers and the partition between water phase and particulate matter has already been occurred mainly in the soil zone, the quality of organic substances in groundwater is dominated by transformation processes. Groundwater systems respond very sensitively on variations of oxygen availability. Continuous changes of the redox-conditions with depth as a result of ongoing oxygen consuming processes (in particular organic matter degradation) influence the constitution of the microbial community and, consequently, the transformation processes affecting organic compounds. The variety of microbially mediated reaction pathways covers

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e.g., nitrate reduction, iron reduction, sulfate reduction and methanogenesis. These processes have high implications in particular for the stability of organic matter in aquifers. Depending on these environmental conditions as well as on the chemical properties of the contaminants either significant microbial degradation but also a stability over long periods of time (years, decades) may be observed.

3.2

Natural Organic Substances

The knowledge on natural substances in ground water is very restricted. Similar to surface water systems a huge proportion of humic substances is proposed (e.g., Alborzfar et al., 1998). Quantitative calculations revealed e.g., amounts of approx. 5 to 20 mg C/L of humic acids in shallow aquifers. However, information on low molecular weight substances are rarely reported. It is known that carboxylic acids contribute to DOC in selected aquifers (McMahon and Chapelle, 1991). Furthermore, some indications for the presence of terpenoids compounds in groundwater and their possible role as humic precursors are reported (Leenheer et al., 2003).

3.3

Anthropogenic Organic Contamination

Anthropogenic pollution is a major concern in particular with respect to shallow aquifers, which frequently represent important drinking water reservoirs. Principally, three major emission sources release organic pollutants into the groundwater systems. Firstly, a direct application of chemicals to soils and their subsequent relocation towards the aquifers contaminates groundwater resources in particular by agrochemicals like pesticides and fertilizers (e.g., Kolpin et al., 2001). Further on, two further types of emission sources are dominating the groundwater contaminations, that release unintentionally pollutants towards the aquifers, sometimes for decades. On the one hand industrial facilities handling with gasoline or petroleum related products, gas production plants and dry-cleaning services are known to have frequently emitted huge amounts of specific pollutants in the past as the result of careless handling or leakages. Typical ground water relevant contaminants related to fossil fuels are monoaromatic compounds especially the BTEX (benzene, toluene, ethylbenzene, xylenes) but also naphthalene and tricyclic aromatic hydrocarbons (e.g., Cozzarelli et al., 1995; Ohlenbusch et al., 2002; Vinzelberg et al., 2005; Zamfirescu and Grathwohl, 2001). Further on, much attention has been given to the gasoline additive methyl tert-butyl ether MTBE, which exhibits a high water solubility and a high environmental stability in groundwater (e.g., Gelmann and Binstock, 2008; Squillace et al., 1996). Dry-cleaning facilities have partially emitted high amounts of the drying agent tetrachloroethylene (PER), what resulted in groundwater contaminations by this compound and its dechlorinated metabolites (e.g., Bradley, 2000; Vieth et al., 2003). Noteworthy, in particular the metabolites exhibit extended persistence under anaerobic conditions. Beside industrial facilities or services a last emissions source of groundwater contamination can be attributed to leakages of waste deposit landfills. Continuous discharge as a result of insufficient bottom sealings has been frequently released a wide spectra of contaminants to the aquifers, because seepage water of deposit landfills are characterized by very complex

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¨ man and Hynning, 1993, Paxeus, 1999, mixtures in particular of organic contaminants (e.g., O Schwarzbauer et al., 2002). Several environmental studies on landfill derived groundwater contamination focused on the distribution and fate of specific organic compounds derived from landfill leachates in the underground (e.g., Albaiges et al., 1986; Heim et al., 2004; Ru¨gge et al., 1995). Intensive activities are related with the remediation of contaminated groundwater either by technical measures or by the so-called natural attenuation approach (e.g., Baun et al., 2003; Eganhouse et al., 2005; Lerner et al., 2005). With respect to the latter remediation approach information on occurrence and rate of microbial degradation is of fundamental importance (Sturchio et al., 1998). Principally two different approaches are used to figure out transformation processes, on the one hand the identification and quantification of metabolites, and on the other hand the monitoring of degradation by compound specific isotope analysis. Examples for characteristic transformations in anaerobic aquifers include the carboxylation of aromatic compounds under sulfate reducing conditions (Coates et al., 2002; Griebler et al., 2004; Meckenstock et al., 2000; Vinzelberg et al., 2005) or the hydroxylation of chlorinated and non-chlorinated aromatics by methanotrophic microbes forming phenolic compounds (e.g., Adriaens and Grbic-Galic, 1994; Coates et al., 2002). Investigations applying isotope analyses have focused mainly on the quantification of degradation processes as a key parameter for natural attenuation approaches (see > Fig. 6). Compound specific analyses have used dominantly stable carbon isotopes but to a minor extend also hydrogen isotopes. A detailed summary of isotope analysis applied to groundwater has been recently published (Schmidt et al., 2004), for principles on isotope analysis see > Chapter 5, Vol. 1, Part 2.

. Figure 6 Spatial distribution of concentration, stable carbon isotope signature, and extent of biodegradation of toluene in a contaminated aquifer (adapted from Griebler et al., 2004).

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4

Marine Environment

4.1

Occurrence and Fate of Organic Compounds in the Marine Environment

Major attention has been attributed to the occurrence and behavior of organic matter in the marine systems since processes causing the natural synthesis as well as degradation or preservation of organic matter in these ecosystems are the initial steps in the generation of kerogen, petroleum and related matter. Hence, the knowledge on the zones of preferential primary production of organic matter by photosynthesis of marine organism as well as the factors affecting the preservation especially in the benthic environment is a main topic in the scientific field of Organic Geochemistry (Killops and Killops, 2005; Tissot and Welte, 1984). Beside the vertical distribution as the result of global ocean currents the dominant process affecting the occurrence of organic matter in the marine environment is its vertical flux within the water column. Thereby, key aspects on the corresponding fate of organic substances are the interaction of dissolved and particulate associated matter in combination with the biological uptake and excretion e.g., by the planctonic food web (Wakeham and Lee, 1989). Principally, organic substances involved in the biological loop (including the bioavailability DOC) are affected by high turn around times, high dynamics and intensive transformation or degradation. However, the adsorption on particulate matter reduces its bioavailability and, therefore, implies an initial step for enhanced preservation. The importance of adsorption phenomena as stabilization processes for labile organic matter was pointed out by Keil et al. (1994) as well as Mayer (1994). Since the reversible adsorption and desorption processes depend among other parameters on the polarity or lipophilicty of the substances to be sorbed, dominantly less polar compounds tend to be more enriched in the solid phase and more effective stabilized by particle association. This particulate matter is partially deposited by sedimentation and the associated, more lipophilic compounds are transferred into the sediments. During sedimentation, comprising a dynamic exchange between POC and DOC by adsorption and desorption, the organic matter undergoes many diagenetic alterations. After its transport through the water column the more anaerobic ambience in the benthic compartment provides further conditions supporting an enhanced environmental stability for many organic compounds accumulated in and incorporated into sediment deposits. The persistence or stability in the benthic and later on in the sedimentary systems over geological times as well as the biotic and abiotic transformation processes affecting the individual molecular structures in this systems are covered by the wide field of Organic Geochemistry and, therefore, is not subject of these chapter. Noteworthy, terrigenous discharge contributes significantly to marine organic matter, in particular at the intersection of marine and terrestrial ecosystems, the estuary and deltas (Hedges et al., 1997). Huge amounts of organic material is discharged from rivers to the coastal regions as dissolved (DOC) but also as particulate organic carbon (POC). Calculation of terrestrial budgets based either on carbon isotope analysis (e.g., Countway et al., 2007; Goni et al., 1997; Raymond and Bauer, 2001; Shi et al., 2001) or determination of indicative chemical marker compounds reflecting unambiguously terrigenous origin e.g., lignin (e.g., Canuel 2001; Harvey and Mannino, 2001; Jaffe et al., 2001; Opsahl et al., 1999). However, calculated budget data vary depending on the approach as well as on the aquatic systems. For the arctic environment a riverine discharge of terrigenous matter to the Arctic Ocean of roughly 25 Tg C yr 1 has been calculated, of which 10 to 40% may reach the Northern Atlantic Ocean (Opsahl et al., 1999). The riverine contribution to the Arctic Ocean is subdivided into

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approx. 80% of DOC and approx. 20% of POC (Dittmar and Kattner, 2003). Global budget calculation on base of lignin analysis suggested an overall contribution of terrigenous material of approximately 0.7–2.4% to the total DOC in the oceans with a predicted shorter oceanic residence time of 20–130 year for this fraction (Opsahl and Benner, 2003).

4.2

Natural Organic Substances

Organic matter in the marine ecosystems originates dominantly from the aquatic organism and, therefore, exhibit partially high similarity to the contributions of biotic organic matter discharged to the terrestrial aquatic systems (see section 1.2.). Beside natural macromolecular like peptides, proteins and polysaccharides (e.g., Khodse et al., 2008) in particular cell membrane lipids and pigments from marine phyto- and zooplankton contributes to the pool of DOC and POC. Well known components comprise n-alkan-1-ols, saturated and unsaturated fatty acids, long chain alkanones (e.g., 37:2 and 37:3 alkenones), di- and tetraphytanyl ethers, steroids (e.g., cholesterol, dinosterol, desmosterol), caretonoids (e.g., lycopane, isorenieratene, diatoxanthin, fucoxanthin, peridinin) and chlorophyll related pigments. According to the general partition behaviour of low molecular weight organic substances as described in Section 1.1, the less polar substances accumulate first in the particulate matter within the water body and, later on, in the sedimentary environment. However, during the passage through the water body high alterations due to biotic and abiotic transformation occur. Examples are autoxidation processes, photoxidation or microbial decomposition (e.g., Rontani et al., 2006; Sun et al., 2004). Further on, marine humic substances contribute dominantly to the DOC, but exhibit structural differences as compared to terrigenous material. A major difference is the degree of aromaticity, which has been already used to discriminate terrestrial and marine humics. Aromatic carbon content has been calculated to be 20–50% in soil humics, 20–35% in peat humics but less than 15% in marine humics. Further on, the H/C atomic ratios vary between 1.0 to 1.5 in marine humics and 0.5 to 1.0 in soil humics (Killops and Killops, 2005). Interestingly, the relative amount of biogenic halogenated compounds is enriched in the marine ecosystems as compared to terrestrial surface water systems probably due to the elevated chloride and bromide concentrations in sea water. These substances cover a wide range of structural diversity including e.g., simple molecules like halogenated methanes and ethanes, but also more complex substances like brominated and chlorinated terpenoics, heterocycles, acetogenins, macrolides and to a higher extend aromatic compounds (Ballschmiter, 2003; Gribbles, 2000). Further compounds, that are still unnoticed so far but are obviously specific marine substances, derived from biomethylation reactions applied to metal ions. One example of such metal organic substances is dimethylthallium, detected in surface water samples from the Atlantic Ocean accompanied by methylated cadmium and lead species (Schedlbauer and Heumann, 2000).

4.3

Anthropogenic Organic Contamination

Many of the environmental aspects of terrestrial contaminants account also for the marine ecosystems due to the discharge of terrestrial matter in coastal areas. Hence, in addition to specific natural compounds serving as terrestrial indicators also anthropogenic contaminants

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are appropriate marker to monitor the riverine impact on the marine environment (Grigoriadou, 2008; Schwarzbauer et al., 2000). Beside river discharge also the aeolian long range transport of particle associated pollutants contributes to the marine contamination. Beside these allochthonous emissions the autothonous immissions are of major interest for the state of pollution of the marine environment. A first aspects is related with shipping activities, which release pollutants during routine operation or as a result of accidents. A common example for the former type of contamination are tin organic compounds, which have been intensively used over a prolonged time as active components in antifouling paints. Noteworthy, not only the marine environment but also rivers, lakes and, in particular, harbours are contaminated by these substances. Most prominent example is tributyl tin, a ionic molecule with lipohilic butyl moieties. As a result of its amphotereous character with respect to its lipo-/hydrophilicity it is accumulated on the one hand in sediments but is also present in high amounts in the water phase. Since, tributyl tin exhibit elevated ecotoxicological effects, it has been substituted recently by other antifouling agents. Shipping activities also result contamination by petroleum derived substances, in particular hydrocarbons. As introduced in Section 1.3, aromatic hydrocarbons can act as indicative substances to characterize petroleum discharge and to differentiate it from combustion derived pollution, which are entering the marine environment via aeolian particles (e.g., Ding et al., 2007). Hence, PAHs has been frequently used (partially together with further petroleum related hydrocarbons like hopanes) to assess the petroleum derived impact on the marine ecosystems, in particular the coastal areas (e.g., Grigoriadou 2008; Yunker and Macdonald, 2003). Beside its marker properties it is also obvious, that PAHs harms the marine environment e.g., by bioaccumulation in marine organisms (Hylland, 2006; Meador et al., 1995, Ohwada et al., 2003). However, PAHs and further petroleum hydrocarbons are constituents of marine sediments not only as the result of shipping activities but also derived from off-shore oil production. For example, drill cuttings has been isolated to contribute significantly to sedimentary hydrocarbon pollution (Scholz-Bo¨tcher et al., 2008, Skaare et al., 2008). A further major source of petroleum related contamination has to be attributed to oil spills. Many accidents of oil tanker have had an enormous impact on the marine ecosystems. The behaviour of oil after release to the marine water depends on various parameters comprising e.g., wind drift with subsequent emulsification and dispersion, the chemcial nature of individual oil fractions, sun light intensity, water temperature and benthic microbial community. Individual fractions of crude oil undergoe different degradation or transformation pathways as well as transport processes. The light components remain over a prolonged time on the water surface and can be subject to abiotic photolysis or photooxidation as well as evaporation. On the contrary, heavier fractions sink to the sea floor. Generally, after oil spills the principal components of oil, the linear alkanes, branched aliphatics, aromatic compounds and functionalised substances, exhibit different residence times on water or in sediments as well as underly varying weathering processes due to their different potential to be microbially degraded (e.g., Ezra et al., 2000; Farias et al., 2008; Gallego et al., 2006). The diverse fate of compound classes has been used to fingerprint oil spills and related sources by biomarkers (e.g., Wang et al., 2006). A high environmental impact has to be associated to the residual fraction of oil remaining in the benthic ecosystems or the beach sediments of the affected coasts over a prolonged time (e.g., Short et al., 2007a). Further long term effects are attributed to the generation of more toxic biotic metabolites or the release of toxic constituents to the sea water after alteration e.g., of the asphaltene fraction (DiToro et al., 2007). Oil spills that have been intensively investigated in terms of long term effects and the distribution and fate of oil

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derived hydrocarbons are, for example, the accidents of the Exxon Valdez in the Gulf of Alaska and the Prestige near the Spanish coast (Bence et al., 1996; Gallego et al., 2006). However, it has to be also mentioned, that a very small proportion of marine petroleum contamination arised from natural seeps discharging oil in particular from coastal regions to the sea (e.g., Short et al., 2007b). In recent years an interesting aspect arised concerning a so far neglected fraction of anthropogenic contaminants, the plastics. Huge amounts of plastic (polypropylene, polystyrene etc.) in different forms (as pellets, foams or foil) have been released to the oceans for decades, although their enormous environmental stability is well known. The ingestion of plastic debris by marine birds and the related harmful effects have been reported since the 80s, whereas the information on ingestion by fishes or filter-feeding organisms are restricted. Also the accumulation of plastic debris in defined regions e.g., of the Pacific Ocean has been elucidated (Moore et al., 2001). Not only the direct harmful impacts also the effects of plastic resin pellets as transport medium for pollutants has been discussed (Mato et al., 2001).

5

Research Needs and Outlook

Although the knowlegde on organic matter in the hydrosphere has been expanded intensively in the last two decades there is still an enormous need to clarify further on the fate of organic substances in water. Interesting aspects of future research are comprising structural elucidation of more complex compounds as well as investigations on the environmental behavior of natural and anthropogenic substances. In particular, our knowledge on natural organic matter in ground water remains on a more or less bulk characterisation. Hence, intensive investigations on the characterisation of dissolved ground water constituents of low molecular as well as macromolecular weight are desirable for the future. Further on, the activities in characterisation of the microbially assisted interaction of metals and organic matter resulting in organo-metal compounds have to be expanded. Further examples for future needful research activities are the investigations on the overall life cycle of natural as well as anthropogenic compounds in the marine systems or the detection and monitoring of disperse or dissolved xenobiotically polymers in river systems.

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Part 5

Biochemistry of Biogenesis

21 Introduction to Microbial Hydrocarbon Production: Bioenergetics M. J. McInerney1 . T. Hoehler 2 . R. P. Gunsalus3 . B. Schink4 Department of Botany and Microbiology, University of Oklahoma, Van Vleet Oval Norman, OK, USA [email protected] 2 Exobiology Branch, NASA Ames Research Center, Moffett Field, CA, USA [email protected] 3 Dept of Microbiology and Molecular Genetics, University of California Los Angeles, Los Angeles, CA, USA [email protected] 4 Department of Biology, University of Konstanz, Konstanz, Germany [email protected]

1

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Introduction: Scope of Microbial Hydrocarbon Production . . . . . . . . . . . . . . . . . . . . . . 322

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Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 324

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Thermodynamics of Microbial Growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 325

4 4.1 4.2 4.3 4.4

Impact of Environmental Conditions on the Thermodynamics of Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 Substrate and Product Concentrations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 327 Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 328 pH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331 Pressure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331

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Thresholds and Minimum Free Energy Change . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 331

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 332

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_21, # Springer-Verlag Berlin Heidelberg, 2010

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Introduction to Microbial Hydrocarbon Production: Bioenergetics

Abstract: Microorganisms play an essential role in the global carbon budget with methanogenesis being a significant global source of methane. The ability to produce hydrocarbons other than methane is widespread among microorganisms and the diversity of hydrocarbon structures that are made is remarkable. However, other than microbial methane production, we know very little about the biochemical processes involved in microbial hydrocarbon formation. Methane production from natural polymers involves a consortium of interacting microbial species. Gibbs free energy yields associated with methanogenesis depend significantly on environmental conditions, especially temperature, activities (concentrations) of substrates and products, and pH, and are typically substantially smaller in natural systems than in growth-optimized cultures. The Gibbs free energy changes involved in the conversion of hydrocarbons, fatty and aromatic acids, alcohols, and hydrogen to methane are close to thermodynamic equilibrium. The low Gibbs free energy changes by which methanogenic consortia operate imply the existence of a minimum free energy change needed to sustain microbial growth, e.g., a biological energy quantum (BEQ), which is support both by theoretical considerations and experimental data. Methanogenic consortia provide excellent models to study interspecies interactions and highly efficient energy economies.

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Introduction: Scope of Microbial Hydrocarbon Production

Microorganisms play a significant role in the production of the simplest hydrocarbon, methane (CH4). Methane is an important fuel and a potent greenhouse gas and its atmospheric concentration has nearly tripled since pre-industrial times (Lelieveld et al., 1998). Estimates for the annual global methane budget range from 500 to 600 Teragram (Tg) (1 Tg equals 1012 g) with about 70% (350–400 Tg) due to microbial activity (Ehhalt et al., 2001). Important sources of microbially produced methane are wetlands including tundra, bogs and swamps, ocean sediments, rice paddies, ruminant animals, oceans, termites, landfills, and waste treatment facilities. Recently, Keppler et al. (2006) showed that plants emit methane, which may account for about 10–30% of methane entering the atmosphere. Because most of the microbially produced methane comes from the decomposition of biomass, methanogenesis is an integral component of the global carbon cycle. Microbial methane production is an ancient process dating to the early Archaean era, 3.5 Gyr ago (Ueno et al., 2006). Microorganisms make a variety of hydrocarbons other than CH4 (> Table 1) (Ladygina et al., 2006; Tornabene, 1980, 1982; Wackett, 2008). Soil microorganisms are significant producers of ethylene (Ilag et al., 1968; Lynch, 1972) and volatile alkanes and alkenes with 2–4 carbons (Ladygina et al., 2006). Geochemical evidence implicates microorganisms in the formation of ethane and propane in deep marine sediments (Hinrichs et al., 2006). Longchain alkane production by marine algae is well documented (> Table 1) although the amounts made by most algae are low (Ladygina et al., 2006; Tornabene 1980, 1982). Brown algae contain n-pentadecane, red algae contain n-heptadecane, and green algae contain C-17cyclopropylalkane (Youngblood and Blumer, 1973). Dunaliella salina produces 6-methylhexadecane and 4-methyl-octadecane (Tornabene, 1980). Cyanbacteria contain C-17-alkanes and methylated alkanes. The green microalga, Botryococcus braunii, is unusual in that it accumulates hydrocarbons up to 75% of its dry mass and may be a promising source for biofuels in the future (Kalacheva et al., 2002). A cobalt-prophryin enzyme was purified from microsomes of B. braunii that decarbonylated octadencanal to heptadecane, CO and some CO2 (Dennis and Kolattukudy, 1992). These data indicate that the pathway for alkane

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. Table 1 Types of hydrocarbons produced by microorganismsa Hydrocarbon type

Microorganisms

CH4

Methanogens

C-2 to C-4 alkanes and alkenes

Many different soil microorganisms

Long-chain alkanes n-Pentadecane (C-15) and n-heptadecane (C-17)

Brown and red algae

C-17 to C-36 alkanes

Botryococcus braunii

Saturated and unsaturated C-17 straight chain hydrocarbons and 6-methyl hexadecane and 4-methyl octadecane

Dunaliella salina (green alga)

Pristane, (C-19), phytane (C-20)

Phototrophic bacteria

C-15 to C-31 alkanes

Various bacteria and fungi

C-17 alkanes; 7, 9-dimethyl hexadecane and 7- and 8-methylheptadecane

Cyanobacteria

Alkenes n-Heneicosahexaene (C-21:6); up to 1% of dry weight

Freshwater and marine algae

Olefins and poly-unsaturated alkenes

Marine algae

Di-unsaturated hydrocarbons; botryococcane

Chlorophytes

C-21 to C-29 alkenes

Micrococcus and Kocuria

Terpenes (Isoprenoids) Isoprene (2-methyl-1,3-butadiene)

Actinomyces, Bacillus subtilis

Carotenes

Fungi, yeasts, algae, bacteria

Squalene (C-30) and isoprenoids, hydroisoprenoids and isopranoids of different chain lengths

Fungi, yeasts, algae, bacteria, and archaea

Lipids Long chain fatty acids (C-12 to C-18)

Bacteria

Mycolic acids (C-60 to C-90)

Mycobacteria, Rhodococcus

Isoprenoids (C-20 to C-40)

Archaea

a

Data from Koga and Mori (2007), Ladygina et al. (2006), Tornabene (1980, 1982), and Wackett (2008) and references therein

synthesize involves the reduction of fatty acids to aldehydes, which are then decarbonylated to alkanes. A number of bacteria are also known to make long-chain alkanes as well as fatty acids (> Table 1). Some members of the genera Micrococcus and Kocuria produce a range of alkenes with 12–29 carbons with subterminal branching. The unsaturated bond is in the middle of the molecule suggesting an interesting biosynthetic reaction possibly involving decarbonylation or decarboxylation and head-to-head condensation of two fatty acids (Tornabene, 1980, 1982). Park (2005) found that membrane fractions of Vibrio furnissi made pentadecane and hexadecane from hexadecanoic acid and detected labeled hexadecanal and hexadecanol from labeled hexadecanoic acid. Pentadecane formation can be explained by the decarbonylation pathway above, but the formation of hexadecane must involve some as yet undescribed mechanism as

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no loss of carbon occurred. However, there is some uncertainty about the ability of V. furnissi to produce large amounts of alkanes (Wackett, 2008). Another important class of hydrocarbons made by microorganisms is terpenes. Actinomyces and Bacillus species are major sources of isoprene (2-methyl-1,3-butadiene) (Ladygina et al., 2006). Mutational analysis shows that isoprene synthesis occurs by the methylerythritol phosphate pathway in Bacillus subtilis (Julsing et al., 2007). Once the intermediates, isopentyldiphosphate and dimethylallyl-diphosphate are formed, terpenes of 10–110 carbons can be made by a series of condensation reactions. Again, details of the pathways are sketchy particularly the mechanism(s) by which carboxylic acid intermediates are converted to hydrocarbons. Finally, all archaea synthesize significant amounts of isoprenoid lipids of C-20 to C40 chain length where considerable variation exists regarding the degree of molecule saturation, cyclization, and methylation (Koga and Morii, 2007). Archaeal isoprenoid biosynthesis proceeds by the mevalonic acid pathway or a modified version of this pathway involving isopentenyl-phosphate rather than diphosphomevalonic acid as an intermediate. Details of these interesting biochemical reactions as well as the microbes involved will be discussed in the chapters subsequent to this section of the handbook.

2

Methanogenesis

The conversion of natural polymers such as polysaccharides, proteins, nucleic acids, and lipids to CO2 and CH4 is called methanogenesis and involves a number of diverse, interacting microbial species. First, numerous fermentative bacteria hydrolyze the polymers and ferment the hydrolysis products to acetate and longer chain fatty acids, CO2, formate, H2 (McInerney et al., 2008; Schink, 1997). Acetogenic bacteria use methanol (from methyl groups of pectin), methyl groups of methoxylated aromatic compounds, some hydroxylated aromatic compounds, and H2 and CO2 to produce acetate (Drake, 1994). A second group of microorganisms works cooperatively with methanogenic bacteria to syntrophically metabolize the products of fermentative metabolism (e.g., propionate and longer chain fatty acids, alcohols, and aromatic acids) to the methanogenic substrates, H2, formate, and acetate. In syntrophic metabolism, the degradation of the parent compound, e.g., the fatty acid, is thermodynamically unfavorable unless the hydrogen, formate and acetate produced by the fatty acid degrader are kept low by the partner methanogens (> Table 2). Lastly, two different groups of methanogens, the hydrogenotrophic methanogens and the acetotrophic methanogens, complete the process by converting the acetate, formate and hydrogen made by other microorganisms to methane and carbon dioxide (Deppenmeier, 2002; Hedderich and Whitman, 2006; Schaefer et al., 1999; Zinder, 1993). The biochemical details of microbial methane production will be discussed in > Chapter 22, Vol. 1, Part 5. In the gastrointestinal tract of animals, only fermentative bacteria and hydrogenotrophic methanogens are active (Hedderich and Whitman, 2006; Mackie and White, 1997). Acetotrophic methanogens and organisms capable of syntrophic metabolism grow too slowly to be effectively maintained in significant numbers in these ecosystems. Thus, organic matter is degraded to acetate and longer chain fatty acids, which accumulate and are absorbed and used by the host animal as energy sources. The amount of energy released per unit of biomass degraded during methanogenesis is very low as most of the energy is retained in methane. For this reason, methanogenesis is the treatment of choice for wastes. Aiyuk et al. (2006) recently reviewed new methanogenic treatment technologies.

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. Table 2 Reactions involved in syntrophic metabolism DGo,a

Reaction Ethanol + H2O ! Acetate + H+ + 2H2

pH2 (atm) for -DG,b

+9.6

equation 61.1 is the Gibbs free energy change of the reaction (e.g., DGr = gD (DG0eD – DG0eA )). Two correlations (61.2 and 61.3 in Heijnen (1999)) are used to calculate D0s /rx if the growth rate is known. The approach is able to predict biomass yields (Ysx, C-mol biomass
C-mol1 of substrate) from the stoichiometry and the thermodynamics of the catabolic reaction so long as a growth rate is known. The approach has an accuracy of about 10–20% over a range of 0.01–0.8 -mol biomass
C-mol1 of substrate for many carbon substrates, but the authors stress that the Ysx values should only be considered a preliminary estimate because there are often more than one biochemical pathway to degraded a compound. Ds01/rx can be considered to be a measure of the amount of biochemical work needed to convert the carbon source into biomass (Heijnen and vanDijken, 1992). Values for Ds01/rx range from 150 to 3,500 kJ (mol biomass C)1. Chemolithotrophic bacteria that use CO2 as a carbon source and use reverse electron transport such as nitrifiers and thiobacilli have high Ds01/rx values. Interestingly, Syntrophobacter fumaroxidans, which needs reverse electron transport for hydrogen production during syntrophic propionate metabolism, also has a very large Ds01/rx value, about 3,500 kJ (mol biomass C)1 (Sholten and Conrad, 2000). The thermodynamic approach predicts that catabolic reactions with more favorable Gibbs free energy changes should result in higher growth yields of the organism and this is generally true. For example, toluene oxidation coupled to aerobic, nitrate and iron respirations (> Table 3) releases large amounts of Gibbs free energy per electron compared to toluene oxidation coupled to sulfate reduction or methanogenesis (> Table 3). Aerobes, denitrifiers and iron reducers that use toluene have higher yields than sulfate reducers or methanogenic consortia (Zwolinski et al., 2000). An interesting question is why, under methanogenic conditions, a consortium is needed to degrade the parent substrate (toluene in this case) to CO2 and CH4, but a single species is able to do so under other electron-accepting conditions. McInerney and Beaty (1988) noted that the Gibbs free energy released per electron for glucose degradation to CO2 and CH4, under methanogenic conditions or to CO2 under sulfatereducing conditions was much lower than that for mineralization of glucose under other electron-accepting conditions or by various glucose fermentations. McCarty (1971) proposed that the free energy released per electron is a major factor determining whether an organism

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will be maintained in anaerobic digestors. Low energy yield per electron should result in low biomass yields according to equation > equation 61.1. Organisms with low cell yields will have difficulties maintaining population size when substrate concentrations are low or when competition for the substrate occurs. While thermodynamic analysis seemingly predicts the appropriate scenario for methanogenic conditions, e.g., a consortium is needed for toluene degradation (Ficker et al., 1999; Meckenstock, 1999), it is not useful in predicting the scenario under sulfate-reducing conditions. Several sulfate reducers completely mineralize toluene in pure culture (Widdel et al., 2006) and interspecies hydrogen transfer was not needed for toluene degradation by aquifer microorganisms under sulfate-reducing conditions (Elshahed and McInerney, 2001). Most likely, considering only the Gibbs free energy released per electron only and without kinetic analyses is too simplistic. Ecological theory predicts that the rate of ATP formation is important. When resources (e.g., substrate) are limiting, organisms that produce ATP at high rates but low yields are favored over those that produce ATP at high yields but at low rates (Pfeiffer et al., 2001). Similarly, kinetic theory for optimal pathway design implies that the optimal pathway length is one that maximizes the rate of ATP production (Costa et al., 2006).

4

Impact of Environmental Conditions on the Thermodynamics of Methanogenesis

The Gibbs free energy yields associated with methanogenesis depend significantly on environmental conditions, and are typically substantially lower in natural systems than in growthoptimized cultures or than is suggested by standard free energy changes. Among factors that influence free energy yields are temperature, activities (concentrations) of substrates and products, pH and pressure when considering their variation in naturally occurring biological systems. The first two have the largest potential effect on energy yield and are considered in some detail below; pH and pressure which have a more modest effect are addressed only briefly.

4.1

Substrate and Product Concentrations

The free energy change of any chemical reaction is affected by variations in the activities of products and reactants, as follows (> 61.2): Q y  P DGr ¼ DG  ðT Þ þ RT  ln Q z ð61:2Þ R where DGr is the free energy available under in situ conditions; DG is the temperatureadjusted free energy change under standard conditions; T is temperature in Kelvin; R is the universal gas constant; PPy and PRx are the mathematical products of the activities of reaction products and reactants, respectively, with each raised to its stoichiometric power (e.g., a reactant having a stoichiometric coefficient of 3 would have its activity raised to the third power in the calculation of DG). For reactions in which environmental activities of substrates and products differ markedly from standard state (defined as 1 M for all species) – especially for species exhibiting high reaction stoichiometry – in situ free energy yields may thus differ dramatically from standard reaction free energy changes.

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Typical environmental substrate and product concentrations result in free energy yields for hydrogenotrophic methanogenesis that are smaller – in some cases by more than an order of magnitude (Hoehler, 2004) – than the standard free energy change (135 kJ·(mol CH4)1 for the reaction written as: CO2 (g) + 4H2 (g) ! CH4 (g) + 3H2O (liq). While the activities of each of the substrates and products in this reaction under environmental conditions are subject to significant natural variation, hydrogen exerts the greatest influence on the energetics of methanogenesis in most systems. This is due to both its high stoichiometric coefficient (and a corresponding 4-fold greater effect than any other species in the reaction) and its typically short half-life (and corresponding potential for rapid change) in many ecosystems (Hoehler et al., 2002). Across a realistic range of hydrogen activities for anaerobic systems in nature, the free energy change of H2-based methanogenesis varies by more than 200 kJ·(mol CH4)1 (> Fig. 1a). In contrast, changes in acetate concentrations yield a much smaller variation in the free energy yield of acetoclastic methanogenesis (CH3COO + H2O ! CH4 + HCO 3 ), by virtue of a unit stoichiometric coefficient (> Fig. 1c). Hydrogen can be delivered to methanogenic communities by either abiotic reactions (e.g., water-rock reactions in hydrothermal systems) or by biological production (e.g., by fermentation of organic matter). The former case may yield H2 activities up to tens to hundreds of millimolar, depending on the fluid source. H2 production by anaerobic fermentation reactions, however, typically encounters thermodynamic inhibition at much lower H2 activities. As with methanogenesis, the high stoichiometric coefficient of H2 in most fermentation reactions + (e.g., fermentation of propionate: CH3CH3COO + 3H2O ! CH3COO + HCO 3 + H + 3H2) renders their energetics highly sensitive to variations in H2 activity (> Fig. 1b). For this reason, H2 concentrations in systems driven by anaerobic decomposition of organic matter are constrained to a range in which both production and consumption of H2 are thermodynamically and biologically favorable (> Fig. 2). The permissible range depends on the specific nature of the fermentation reactions, as well as substrate and product activities and temperature (see below), but this phenomenon generally results in methanogenic energy yields much lower than are typical of growth-optimized cultures. A comparable effect can also be inferred for acetate production/consumption (> Figs. 1d, c). Notably, however, the freeenergy changes of both fermentative production and methanogenic consumption of acetate are four-fold less sensitive to acetate concentrations than in the corresponding case of H2. Free energy yields for acetoclastic methanogenesis are therefore constrained within a narrower range across the spectrum of environmentally-realistic acetate concentrations, and differences between natural systems and cultures are less exaggerated than for H2-based methanogenesis.

4.2

Temperature

Methanogens are represented across much of the biologically tolerated range of temperature, and variation over this range can significantly impact the free energy yield of methanogenesis. Temperature has two main effects on reaction energetics, as indicated by > equation 61.2. First, the entropic contribution to free energy increases linearly with temperature, DG = DH –TDS, so that the standard free energy change, DG , is temperature-sensitive. For reactions having large entropy changes, this effect can be quite pronounced across the biologically tolerated range of temperatures, especially when considering processes that occur with small in situ free energy yields. This is the case for H2-consuming methanogenesis and is generally true for most fermentation reactions that involve high stoichiometries of H2 production. Second,

. Figure 1 The effect of activity and temperature on the Gibbs free energy change for: (a) hydrogenotrophic methanogenesis versus aqueous H2 activity; (b) syntrophic propionate metabolism versus aqueous H2 activity; (c) acetoclastic methanogenesis aqueous acetate activity; and (d) syntrophic propionate metabolism versus aqueous acetate activity. Values are plotted for four temperatures: 0 C (solid line), 25 C (long dashes), 50 C (short dashes), and 100 C (alternating short/long dashes). Reaction free energy changes were calculated using Equation 61.3. Standard free energies of reaction were calculated using thermodynamic data from Shock and Helgeson (1988, 1990), with temperature dependence calculated via the Gibbs-Helmholtz equation (for the latter calculation, the small temperature dependence of DH over the range considered was neglected). Calculations are for the reactions shown, with all species considered in the aqueous form. The standard free energy change for hydrogenotrophic methanogenesis (written as HCO3 (aq) + 4 H2 (aq) + H+ (aq) ! CH4 (aq) + 3H2O (liq)) is -229 kJ·(mol CH4)1. Calculations assume the following concentrations (M), unless otherwise indicated by a variable axis: propionate and acetate = 10–5; HCO3 = 2·10–2; CH4 = 10–3; H+ = 10–7; H2 = 10–7.

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. Figure 2 The zone where the production and consumption of H2 are both thermodynamically favourable. The shade region represents the range of H2 activities at 25 C in which hydrogenotrophic methanogenesis and syntrophic propionate fermentation are thermodynamically favorable (note that this range may be smaller when considering biological minimum free energy thresholds). Syntrophic conversion of propionate to methane is not possible outside of this range. The open symbol labeled ‘‘25 C’’ represents the H2 activity at which the free energy yields for the methanogenic and syntrophic reactions are equalized, along with the energy yield associated with each reaction. Equivalent values are also plotted as open symbols for 0, 50, and 100 C. Note that the mid-point H2 activity increases by more than 3 orders of magnitude over this range in temperature, while the free energy yield available to each reaction increases slightly.

the impact of substrate and product activities that deviate from standard (unit activity) conditions becomes more pronounced with increasing temperature. This effect is most important for reactions in which one or more species have in situ activities that lie far from unit activity, particularly if those species have high stoichiometric coefficients, e.g., syntrophic acetate metabolism where 4 H2 are made per acetate. Again, this is generally the case for naturally occurring H2-consuming methanogenesis and fermentation reactions that produce H2 with high stoichiometry, while the effect is less pronounced for acetoclastic methanogenesis. The effects of temperature on free energy yields of methanogenesis and propionate fermentation are shown in > Fig. 1. Noting that this magnitude is (by virtue of the second temperature effect) specific to the choice of substrate and product activities, it is nonetheless clear that, for environmentally plausible conditions, temperature shifts across the biologically tolerated range can cause free energy yields to increase or decrease by several times the typical in situ yield. Such changes (particularly in the positive direction) may significantly shift the range of substrate or product concentrations over which a given reaction is favorable. As shown in > Fig. 2, for example, the range of H2 activities over which both methanogenesis and propionate fermentation are favourable (and the mid-point H2 activity, at which the energy yields of the two reactions are equal) shifts by more than three orders of magnitude in going from 0 to 100 C.

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4.3

21

pH

The effects of environmental pH on free energy yield, though potentially substantial if calculated over the full range of pH in natural aquatic systems, are less straightforward to interpret. These effects can be direct, for reactions that involve ‘‘H+’’ or ‘‘OH’’ as substrates or products, or indirect, by affecting the speciation of pH-sensitive products or reactants (e.g., carbonates, organic acids, etc.). However, given extremes of pH, organisms can employ a variety of mechanisms to regulate the pH of the intracellular medium (where biochemical reaction energetics must be calculated) at more clement levels than the surrounding environment (Krulwich, 1995, 2000). The specific mechanism employed, and extent to which pH is regulated, may thus variably mitigate and modify impacts on free energy change that are predicted based on environmental pH values.

4.4

Pressure

Pressure exhibits a natural range of 3–4 orders of magnitude across biological systems, from atmospheric (approx. 1 bar) to >1,000 bar in ocean trench or deep subsurface ecosystems. However, the direct effect of this range on the thermodynamics of aqueous reactions (including the metabolic reactions of methanogens) is modest. For example, holding all other factors constant, a pressure change from 1 to 1,000 bar changes the free energy change associated with hydrogenotrophic methanogenesis changes by only about 3 kJ·(mol CH4)1. The secondary, and potentially more important, effect of pressure in biological systems is on the solubility of gaseous substrates or end-products where H2 and CH4 can (and do) reach dissolved concentrations hundreds of times higher at the ocean floor than in surface ecosystems (Boetius et al., 2000). Because it is the dissolved concentration of substrate or product that effects the free energy changes of aqueous biochemical reactions, this effect can significantly alter the energetics of methanogenesis. The impact of changing substrate and product concentrations is considered below. In summary, the impacts of temperature, pH, pressure, and non-standard activities of substrates and products, collectively, can shift in situ free energy yields dramatically away from values suggested by standard free energies of reaction, or from those typically experienced by organisms grown in culture. This effect is particularly pronounced for processes that produce or consume H2 with high stoichiometry, such as hydrogenotrophic methanogenesis and many fermentation reactions.

5

Thresholds and Minimum Free Energy Change

As discussed above (> equation 61.2), there is a strong effect of in situ concentration on the Gibbs free energy of the catabolic reaction for some anaerobic processes. The activity of methanogens is essential to maintain hydrogen and formate levels low enough for syntrophic metabolism to be energetically favorable (> Table 2; > Figs. 1 and 2) (Schink, 1997). Consistent with thermodynamic predictions, thresholds for substrate metabolism, defined as the concentration below which further substrate decay is not observed, occur for syntrophic metabolism and methanogenesis (Cord-Ruwisch et al., 1988; Dwyer et al., 1988; Hoehler, 2004;

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Jackson and McInerney, 2002; Lovley, 1985; Scho¨cke and Schink, 1997; Seitz et al., 1990; Warikoo et al., 1996). The free energy changes for syntrophic metabolism and methanogenesis approach a minimum free energy value when substrate thresholds are reached (Hoehler, 2004; Schink, 1997). This minimum free energy, also referred to as the biological energy quantum (BEQ), predicts the favorability of continuous biological activity. BEQ values 12–15 kJ mol1 of substrate have been estimated based on the free energy change needed for ATP synthesis under physiological conditions (about 60 to 70 kJ mol1) and the number of protons needed to make an ATP molecule by ATP synthase (3 to 5 proton per ATP) (Hoehler, 2004; Schink, 1997; Schink and Stams, 2002). Experimental evidence shows that several syntrophic metabolisms operate at free energy changes in the range of 15–20 kJ mol1, close to the theoretically predicted BEQ (Schink, 1997; Scholten and Conrad, 2000). The small amounts of free energy released during syntrophic metabolism must be shared among the partners (Schink, 1997). Thus, it is appropriate to describe syntrophy as an extreme existence, a lifestyle that involves a marginal or equilibrium energy economy, where the direction of metabolism depends on the prevailing environmental conditions. The existence of an equilibrium energy economy was recently illustrated by the metabolism of Thermacetogenium phaeum (Hattori et al., 2005). T. phaeum is a fascinating microorganism that syntrophically oxidizes acetate to CO2 and H2 in coculture with a methanogen where the hydrogen partial pressure is low. It also makes acetate from CO2 and H2 in pure culture where the hydrogen partial pressure is high. The enzymes of the Wood-Ljungdahl pathway are present under both acetate utilization and acetate formation conditions (Hattori et al., 2005). Interestingly, cocultures containing T. phaeum immediately switch from syntrophic acetate oxidation to homoacetogenic acetate formation depending on the environmental conditions. An important unresolved question is how T. phaeum manages to make ATP if it can switch its metabolism so easily. There must be unique steps linked to ATP formation in one direction that are decoupled from ATP hydrolysis in the reverse direction. This could possibly involve some type of switch in electron flow in the membrane, but the mechanisms for this has not yet been identified. The complete sequencing of the genomes of several bacteria capable of syntrophic metabolism is underway and may provide insights into the hallmarks of their interesting physiology.

6

Research Needs

The global importance of microbial methane production is well documented; however, we do not fully understand the importance or function of the microbial production of many hydrocarbons other than methane. Hydrocarbon production occurs among diverse microorganisms. In many cases, the amount of hydrocarbons produced is low and the biochemical mechanisms for their formation are poorly understood. In contrast, a diverse range of fatty acids and isoprenoid lipids are synthesized for incorporation into bacterial and archaeal cell membranes. However many details of their pathways and/or metabolic control are poorly understood. Clearly, much work is needed if we are to exploit non-methane microbial hydrocarbon metabolism for biofuel production. Microbial hydrocarbon metabolism may provide a means for the continued use of hydrocarbons in a carbon-neutral fashion. Methanogenesis and in particular syntrophic metabolism operate at very small free energy changes, suggesting a minimum free energy change is needed to sustain biological activity.

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However, the biochemical mechanisms of energy conservation and its regulation in bacteria capable of syntrophic metabolism are poorly understood. The stoichiometry of ions translocated per mole substrate consumed by the syntrophic metabolizer in addition to the stoichiometry of ions consumed in support of ATP synthesis are critical issues that remain unresolved. Bacteria and archaea reside at the interface between the inhabited and uninhabited realms of our planet. They represent the ultimate biological arbiters of chemical exchange between those spheres. In some environments such as the deep subsurface, the energy flux and growth rates are orders of magnitude below anything we have observed in the laboratory. How is it possible to maintain complex microbial communities and critical cell functions at energy economies that barely allow cell growth? Do these organisms have properties beyond our current understanding of microbial biochemistry, or are energy sources available that we have not yet identified?

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for the Microbial Community, M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.). 3rd edn. New York: Springer-Verlag, pp. 1050–1079. Heider J, Spormann AM, Beller HR, Widdel F (1999) Anaerobic bacterial metabolism of hydrocarbons. FEMS Microbiol Rev 22: 459–473. Heijnen JJ (1999) MC Flickinger, MCSW Derew (eds.). SWEncyclopedia of Bioprocess Technology: Fermentation, Biocatalysis, and Bioseparation. New York: Wiley, pp. 267–291. Heijnen JJ, vanDijken JP (1992) In search of a thermodynamic description of biomass yields for the chemotrophic growth of microorganisms. Biotechnol Bioeng 39: 833–858. Hinrichs K-U, Hayes JM, Bach W, Spivack AJ, Hmelo LR, Holm NG, Johnson CG, Sylva SP (2006) Biological formation of ethane and propane in the deep marine subsurface. Proc Natl Acad Sci USA 103: 14684–14689. Hoehler T (2004) Biological energy requirements as quantitative boundary conditions for life in the subsurface. Geobiology 2: 205–215. Hoehler TM, Albert DB, Alperin MJ, Bebout BM, Martens CS, DesMarais DJ (2002) Comparative ecology of H2 cycling in sedimentary and phototrophic ecosystems. Antonie van Leeuwenhoek 81: 575–585 Ilag L, Curtis RW (1968) Production of ethylene by fungi. Science 159: 1357–1358. Jackson BE, McInerney MJ (2002) Anaerobic microbial metabolism can proceed close to thermodynamic limits. Nature 415: 454–456. Jones DM, Head IM, Gray ND, Adams JJ, Rowan AK, Aitken CM, Bennett B, Huang H, Brown A, Bowler BFJ, Oldenburg T, Erdmann M, Larter SR (2008) Crude oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 451: 176–180. Julsing MK, Rijpkema M, Woerdenbag HJ, Quax WJ, Kayser O (2007) Functional analysis of genes involved in biosynthesis of isoprene in Bacillus subtilis. Appl Microbiol Biotechnol 75: 1377–1384. Kalacheva GS, Zhila NO, Volova TG (2002) Lipid and hydrocarbon compositions of a collection strain and a wild sample of green microalga Botryococcus. Aquat Ecol 36: 317–330. Keppler F, Hamilton JTG, Braß M, Ro¨ckmann T (2006) Methane emissions from terrestrial plants under aerobic conditions. Nature 439: 187–191. Koga Y, Morii H (2007) Biosynthesis of ether-type polar lipids in Archaea and evolutionary considerations. Microbiol Mol Biol Rev 71: 97–120. Krulwich TA (1995) Alkaliphiles: ‘basic’ molecular problems of pH tolerance and bioenergetics. Mol Microbiol 15: 403–410. Krulwich TA (2000) Alkaliphilic Prokaryotes. In The Prokaryotes: An Evolving Electronic Resource for the

Microbial Community, vol. 2. M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.). New York: Springer-Verlag, pp. 309–336 Ladygina N, Dedyukhina EG, Vainshtein MB (2006) A review on microbial synthesis of hydrocarbons. Process Biochem 41: 1001–1014. Lelieveld J, Crutzen PJ, Dentener FJ (1998) Changing concentration, lifetimes and climate forcing of atmospheric methane. Tellus B 50: 128–150. Lovley DR (1985) Minimum threshold for hydrogen metabolism in methanogenic bacteria. Appl Environ Microbiol 49: 1530–1531. Lynch JM (1972) Identification of substrates and isolation of microorganisms responsible for ethylene production in the soil. Nature 240: 45–46. Mackie RI, White BA (eds.). (1997) Gastrointestinal microbiology, vol. 1. New York: Chapman & Hall. McCarty PL (1971) Energetics and kinetics of anaerobic treatment. In Anaerobic Biological Treatment Processes. RF Gould (ed.). Washington, DC: American Chemical Society, pp. 91–107. McInerney MJ, Beaty PS (1988) Anaerobic community structure from a nonequilibrium thermodynamic perspective. Can J Microbiol 34: 487–493. McInerney MJ, Struchtemeyer CG, Sieber J, Mouttaki H, Stams AJM, Schink B, Rohlin L, Gunsalus RP (2008) Physiology, ecology, phylogeny and genomics of microorganisms capable of syntrophic metabolism. Ann N Y Acad Sci 1125: 58–72. Meckenstock RU (1999) Fermentative toluene degradation in anaerobic defined syntrophic cocultures. FEMS Microbiol Lett 177: 67–73. Park M-O (2005) New pathway for long-chain n-alkane synthsis via 1-alcohol in Vibrio furnissii M1. J Bacteriol 187: 1426–1429. Pfeiffer T, Schuster S, Bonhoeffer S (2001) Cooperation and competition in the evolution of ATP-producing pathways. Science 292: 504–507. Schaefer G, Engelhard M, Mueller V (1999) Bioenergetics of the Archaea. Microbiol Mol Biol Rev 63: 570–620. Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61: 262–280. Schink B, Stams AJM (2002) Syntrophism among prokaryotes. In The Prokaryotes: An Evolving Electronic Resource for the Microbial Community, M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.). New York: Springer-Verlag, pp. 309–336. Scho¨cke L, Schink B (1997) Energetics of methanogenic benzoate degradation by Syntrophus gentianae in syntrophic coculture. Microbiology 143: 2345–2351. Scholten JC, Conrad R (2000) Energetics of syntrophic propionate oxidation in defined batch and chemostat cocultures. Appl Environ Microbiol 66: 2934–2942.

Introduction to Microbial Hydrocarbon Production: Bioenergetics Seitz HJ, Schink B, Pfennig P, Conrad R (1990) Energetics of syntrophic ethanol oxidation in defined chemostat cocultures. 1. Energy requirement for hydrogen production and hydrogen oxidation. Arch Microbiol 155: 82–88. Shock EL, Helgeson HC (1988) Calculation of the thermodynamic and transport properties of aqueous species at high pressures and temperatures: correlation algorithms for ionic species and equation of state predictions to 5 kb and 1000 C. Geochim Cosmochim Acta 52: 2009–2036. Shock EL, Helgeson HC (1990) Calculation of the thermodynamic and transport properties of aqueous species at high pressures and temperatures: standard partial molal properties of organic species. Geochim Cosmochim Acta 54: 915–945. Thauer RK, Jungermann K, Decker K (1977) Energy conservation in chemotrophic anaerobic bacteria. Bacteriol Rev 41: 100–180. Tornabene TG (1980) Formation of hydrocarbons by bacteria and algae. Basic Life Sci 18: 421–438. Tornabene TG (1982) Microorganisms as hydrocarbon producers. Experientia 38: 43–46. Ueno Y, Yamada K, Yoshida N, Maruyama S, Isozaki Y (2006) Evidence from fluid inclusions for microbial methanogenesis in the early Archaean era. Nature 440: 516–519.

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Wackett LP (2008) Microbial-based motor fuels: science and technology. Microbial Biotechnol 1: 211–225. Warikoo V, McInerney MJ, Robinson JA, Suflita JM (1996) Interspecies acetate transfer influences the extent of anaerobic benzoate degradation by syntrophic consortia. Appl Environ Microbiol 62: 26–32. Widdel F, Boetius A, Rabus R (2006) Anaerobic biodegradation of hydrocarbons including methane. In The Prokaryotes: An Evolving Electronic Resource for the Microbial Community, M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.). New York: Springer-Verlag, pp. 1028–1049. Wilhelms A, Larter SR, Head I, Farrimond P, di-Primio R, Zwach C (2001) Biodegradation of oil in uplifted basins prevented by deep-burial sterilization. Nature 411: 1034–1037. Youngblood WW, Blumer M (1973) Alkanes and alkenes in marine benthic algae. Marine Biol 21: 163–172. Zinder SH (1993) Physiological Ecology of Methanogens. In Methanogenesis; Ecology, Physiology, Biochemistry & Genetics. JG Ferry (ed.). London: Chapman & Hall, pp. 128–206. Zwolinski MD, Harris RF, Hickey WJ (2000) Microbial consortia involved in the anaerobic degradation of hydrocarbons. Biodegradation 11: 141–158.

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22 Methanogenesis: Syntrophic Metabolism J. R. Sieber1 . M. J. McInerney1,* . C. M. Plugge2 . B. Schink3 . R. P. Gunsalus4 1 Department of Botany and Microbiology, University of Oklahoma, Norman, OK, USA [email protected] *[email protected] 2 Laboratory of Microbiology, Wageningen University, Wageningen, The Netherlands [email protected] 3 Department of Biology, Universita¨t Konstanz, Konstanz, Germany [email protected] 4 Department of Microbiology and Molecular Genetics, Los Angeles, CA, USA [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 338

2

Importance of Syntrophy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 338

3

Bioenergetic Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 339

4

Interspecies Electron Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 340

5 5.1 5.2 5.3 5.4

Biochemical Pathways for Syntrophic Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341 Acetate Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 341 Propionate Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 343 Butyrate Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 345 Benzoate Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 346

6

Potential Mechanisms for Reverse Electron Transport . . . . . . . . . . . . . . . . . . . . . . . . . . . . 348

7

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 350

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_22, # Springer-Verlag Berlin Heidelberg, 2010

338

22

Methanogenesis: Syntrophic Metabolism

Abstract: Syntrophy is a mutualistic interaction in which two metabolically different types of microorganisms are linked by the need to keep metabolites exchanged between the two partners at low concentrations to make the overall metabolism of both organisms feasible. In most cases, the cooperation is based on the transfer of hydrogen, formate, or acetate from fermentative bacteria to methanogens to make the degradation of electron-rich substrates thermodynamically favorable. Syntrophic metabolism proceeds at very low Gibbs’ free energy changes, close to the minimum free energy change needed to conserve energy biologically, which is the energy needed to transport one proton across the cytoplasmic membrane. Pathways for syntrophic degradation of fatty acids predict the net synthesis of about onethird of an ATP per round of catabolism. Syntrophic metabolism entails critical oxidationreduction reactions in which hydrogen or formate production would be thermodynamically unfavorable unless energy is invested. The membrane processes involved in ion translocation and reverse electron transport are poorly understood. While much evidence supports interspecies transfer of hydrogen and formate, other mechanisms of interspecies electron transfer exist including cysteine cycling and possibly direct electron transfer by electrically conductive pili.

1

Introduction

Syntrophy is an energetically limited interaction between cells of different species, e.g., the fatty acid degrader and the methanogen (> Table 1) (McInerney et al., 2008; Schink, 1997; Schink and Stams, 2006). The mutual dependence between the two metabolic types of organisms is so extreme that neither one functions without the activity of its partner. Together, the partners perform functions that neither one can do alone. The degradation of the respective substrate, in this case, a fatty or aromatic acid (> Table 1), is thermodynamically unfavorable if the product concentrations are at standard conditions (1 M concentration, or 1 atm for gases). The function of methanogens is to consume hydrogen, for example, to low steady-state pressure (10–4–10–5 atm) to make fatty and aromatic acid oxidation thermodynamically favorable (> Table 1).

2

Importance of Syntrophy

Syntrophic metabolism is an essential, but the least energetically favorable, step in the conversion of organic matter to methane and carbon dioxide in anoxic environments. Biological methane production, also termed methanogenesis, is an important process in the global carbon cycle, accounting for about 1–2% of the carbon fixed annually by photosynthesis (Hedderich and Whitman, 2006). Annual global methane emissions into the atmosphere are large, about 500–600 Teragram (Tg) (1 Tg equals 1012 g), and more than 70% (350–400 Tg) of these emissions are due to microbial activity (Ehhalt et al., 2001). Syntrophic metabolism is often the rate-limiting step in methanogenesis (McCarty, 1971; McInerney et al., 1981) and, thus, is an important process controlling the global carbon flux. The degradation of natural polymers such as polysaccharides, proteins, nucleic acids, and lipids to CO2 and CH4 involves a complex microbial community (McInerney et al., 1981; Schink and Friedrich, 1994). Fermentative bacteria hydrolyze the polymeric substrates such as polysaccharides, proteins, and lipids, and ferment the hydrolysis products to acetate, longerchain fatty acids, CO2, formate, and H2. Propionate and longer-chain fatty acids, alcohols, and

22

Methanogenesis: Syntrophic Metabolism

some amino acids and aromatic compounds are syntrophically metabolized to the methanogenic substrates: H2, formate, and acetate (Schink, 1997; Schink and Stams, 2006). Lastly, two different groups of methanogens, the hydrogenotrophic methanogens and the acetotrophic methanogens, complete the process, converting acetate, formate and hydrogen produced by other microorganisms to methane and carbon dioxide. The syntrophic degradation of fatty and aromatic acids accounts for much of the carbon flux in methanogenic environments (McCarty, 1971; Pavlostathis and Giraldo-Gomez, 1991). Initial anaerobic transformations of aromatic compounds (Heider and Fuchs, 1997a, b; Schink et al., 2000) generally lead to the conversion of diverse aromatic compounds into benzoylcoenzyme A (CoA) (Breese and Fuchs, 1998; Gallert and Winter, 1994; Gibson et al., 1994, 1997; Hirsch et al., 1998; Merkel et al., 1989). In methanogenic environments, the reduction and cleavage of the aromatic ring are catalyzed by syntrophic associations of benzoatedegrading microorganism and hydrogen- and/or formate-using methanogens (Ferry and Wolfe, 1976; Mountfort and Bryant, 1982; Szewzyk and Schink, 1989).

3

Bioenergetic Considerations

Syntrophy is a fascinating process from a bioenergetic perspective. Even if methanogenic activity is high and hydrogen levels are low, syntrophic metabolism releases very little free energy, which must be shared among the partners (Schink, 1997). Organisms capable of syntrophic metabolism operate at free energy changes very close to the minimum increment of energy required for ATP synthesis (Hoehler, 2004; Schink, 1997). The minimum amount of energy needed for ATP synthesis is predicted to be about
23 kJ mol
1 although this value maybe as low as
15 to
20 kJ mol
1. Most of the free energy changes observed during syntrophic metabolism are in this range (Schink, 1997) although some studies have found free

. Table 1 Reactions involved in syntrophic metabolism 0

Reactions

0

DG0 a (kJ/mol)

DG b (kJ/mol)


135.6


15.8


130.4


11.8

+104.6


1.5

+76.1


16.9

+48.6


39.2

+70.1


68.5

Methanogenic reactions 4H2 + HCO3– + H+ ! CH4 + 3H2O 4HCOO + H2O + H ! CH4 + –

+

3HCO
3

Syntrophic oxidations Acetate– + 4H2O ! 2HCO3– + H+ + 4H2 Propionate + 3H2O ! Acetate + –

–

HCO
3 +

+ H + 3H2

HCO
3

+

+

Butyrate– + 2H2O ! 2 Acetate– + H + 2H2 Benzoate + 7H2O ! 3 Acetate + –

a

–

+ 3H + 3H2

Calculated from the data in Thauer et al. (1977) with the free energy of formation for benzoate given in Kaiser and Hanselmann (1982) b Calculated on the basis of the following conditions observed in methanogenic ecosystems: partial pressures of H2 of 1 Pa and of CH4 of 50 kPa, 50 mM bicarbonate, and the concentrations of the substrates and acetate at 0.1 mM each

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energy changes less than
10 kJ mol
1 (Dwyer et al., 1988; Scholten and Conrad, 2000). An important question is how these organisms exploit such small free energy changes for growth. The second fascinating feature of syntrophic metabolism is the need for reverse electron transport. In syntrophic metabolism, there are critical oxidation-reduction reactions that are thermodynamically unfavorable, e.g., result in a negative DE0 . For example, the production of hydrogen (E0 of
261 mV at 1 Pa H2) or formate (E0 of
258 mV at 1 mM formate) (Schink, 1997) from electrons generated from the oxidation of acyl-CoA intermediates to their respective enoyl-CoA intermediates (E0 of
10 mV) (Sato et al., 1999) has a DE0 of about
250 mV. A hydrogen partial pressure of about 10–5 Pa would make this reaction thermodynamically favorable (Schink, 1997). The syntrophic metabolism of propionate by Syntrophobacter wolinii by the methylmalonyl-CoA pathway (Houwen et al., 1990) involves the oxidation of succinate to fumarate (E00 of + 33 mV) (Thauer et al., 1977). Here again, a very low hydrogen partial pressure (10–6 Pa) is needed for hydrogen production to be thermodynamically favorable (Schink, 1997). Methanogens cannot generate such low hydrogen partial pressures because hydrogenotrophic methanogenesis reaches thermodynamic equilibrium at 0.2 Pa H2. Hydrogen or formate production can occur only with energy input, a process called reverse electron transport. The most likely energy source for this energy input is an ion gradient. Consistent with the requirement for an ion gradient for hydrogen production, the protonophore (CCCP) and the ATP synthase inhibitor (DCCD) inhibited hydrogen production from butyrate by Syntrophomonas wolfei and from benzoate by Syntrophus buswellii (Wallrabenstein and Schink, 1994). Similarly, hydrogen formation from glycolate by membrane vesicles of Syntrophobotulus glycolicus (Friedrich et al., 1996) required ATP or a proton gradient (Friedrich and Schink, 1993, 1995). While it is clear that reverse electron transport is needed for syntrophic metabolism, the nature of this system is not known. How do syntrophic microbial associations operate at low energy conditions? Do they have novel mechanisms for energy conservation or are they more efficient at conserving energy than other microorganisms? We will analyze what is known about syntrophic metabolism in an attempt to answer these questions. Further details on the physiology of the organisms capable of syntrophic metabolism are available in several comprehensive reviews (McInerney et al., 2008; Schink, 1997; Schink and Stams, 2006).

4

Interspecies Electron Transfer

Above, we defined syntrophy based on the exchange of H2 between the syntrophic partners. However, other mechanisms to exchange electrons equivalents exist. Most hydrogenotrophic methanogens use formate or hydrogen or both (Hedderich and Whitman, 2006; Liu and Whitman, 2008). There is very little difference in free energy change for methane production when hydrogen versus formate serves as the electron donor (> Table 1). The conclusion of many studies is that syntrophic metabolism can involve either interspecies transfer of hydrogen and/or formate. Syntrophic metabolism by hydrogen transfer was shown for glycolate metabolism by Syntrophobotulus glycolicus (Friedrich et al., 1996), sugar metabolism by Syntrophococcus sucromutans (Krumholz and Bryant, 1986), acetate metabolism by a thermophilic, syntrophic acetate-oxidizing strain AOR (Lee and Zinder, 1988b) and ethanol metabolism by S-organism (Bryant et al., 1967), by culturing these organisms with a methanogen that uses only hydrogen. In a similar fashion, syntrophic metabolism by formate transfer was shown for an amino acid degrader by using a sulfate-reducing partner that uses formate but

Methanogenesis: Syntrophic Metabolism

22

not H2 (Zindel et al., 1988). Syntrophic propionate degradation by Syntrophobacter fumaroxidans (Dong et al., 1994b; Dong and Stams, 1995) and syntrophic butyrate degradation by Syntrophomonas (Syntrophospora) bryantii (Dong et al., 1994a) occurred only with a methanogen that used both hydrogen and formate, and not with a methanogen that used only hydrogen, implicating the need for formate metabolism. Proteomic and enzymatic analyses showed high levels of expression of formate dehydrogenase in both S. fumaroxidans and its methanogenic partner, arguing for the importance of formate metabolism (de Bok et al., 2002a, b, 2003). Flux analysis of this coculture (de Bok et al., 2002a) and of a butyratedegrading coculture (Boone et al., 1989) indicated that the H2 diffusion was too slow to account for the rates of syntrophic metabolism. The use of hydrogen and/or formate as the interspecies electron carrier provides an explanation why so many methanogens use both hydrogen and formate. Genomic analysis supports the involvement of both compounds because the genomes of Syntrophus aciditrophicus, Syntrophomonas wolfei and Methanospirillum hungatei, the methanogenic partner most often observed in syntrophic associations, have multiple formate dehydrogenase and hydrogenase genes (McInerney et al., 2007; Sieber et al., 2008). Molecules other than hydrogen or formate may be involved in interspecies electron transfer and their use may circumvent the need for reverse electron transport. An acetateoxidizing coculture of Geobacter sulfurreducens and Wolinella succinogenes used cysteine as the interspecies electron carrier (Kaden et al., 2002). Cysteine is commonly used as a reductant in anaerobic media, which could explain why it was overlooked as an electron carrier. Another possibility is direct electron transfer between the syntrophic partners by electron conductive pili or nanowires (Gorby et al., 2006; Reguera et al., 2005). Interspecies electron transfer by nanowires is difficult to prove in syntrophic associations because we do not have the ability to mutate the pilus genes in either of the syntrophic partners at this time. However, nanowire-like structures connecting the syntrophic propionate degrader, Pelotomaculum thermopropionicum with its methanogenic partner have been observed by electron microscopy (Gorby et al., 2006; Ishii et al., 2005) and scanning tunneling microscopy showed that these structures were electron transmissive (Gorby et al., 2006). Some researchers point to aggregation of cells in cocultures as proof of direct electron transfer (Logan and Regan, 2006), but aggregation also reduces the distance between the syntrophic partners, which would increase the rate of hydrogen or formate transfer (Conrad and Zeikus, 1985; Ishii et al., 2005; Thiele and Zeikus, 1988).

5

Biochemical Pathways for Syntrophic Metabolism

The pathways for several syntrophic metabolisms are known and an analysis of the bioenergetics of these pathways illustrates how small amounts of energy are conserved during syntrophic metabolism (Schink, 1997).

5.1

Acetate Metabolism

Syntrophic acetate metabolism is a remarkable process that supports the concept that syntrophic metabolism is very energy efficient. A thermophilic organism, strain AOR, and Thermacetogenium phaeum produce acetate when grown axenically with H2 and CO2, and oxidize acetate when grown syntrophically (Hattori et al., 2001; Lee and Zinder, 1988a). These organisms apparently use the same pathway, the Wood–Ljungdahl pathway, for both

341

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acetate synthesis and its oxidation (> Fig. 1). Several of the key enzymes of this pathway (acetyl-CoA synthase/carbon monoxide dehydrogenase and formate dehydrogenase) were detected under each growth condition (Hattori et al., 2005; Lee and Zinder, 1988a). Both organisms partner with methanogens capable of using formate and hydrogen, although strain AOR couples better with methanogens that only use hydrogen (Lee and Zinder, 1988b). When T. phaeum was grown in a methanogenic coculture, it was able to immediately switch from syntrophic acetate oxidation to homoacetogenic acetate formation indicating that the entire enzyme apparatus operates in a reversible manner (Hattori et al., 2005). Reversibility of metabolism suggests near-equilibrium conditions and high efficiency of energy conservation. Other organisms capable of syntrophic acetate metabolism include Geobacter sulfurreducens (Cord-Ruwisch et al., 1998), ‘‘Candidatus Contubernalis alkalaceticum’’ (Zhilina et al., 2005), Clostridium ultunense (Schnu¨rer et al., 1997), and Thermotoga lettingae (Balk et al., 2002). G. sulfurreducens metabolizes acetate through the citric acid cycle (Galushko and

. Figure 1 Pathway for acetate oxidation and synthesis in syntrophic acetate oxidizers, adapted from Hattori (2008). The enzymes involved are as follows: (1) acetate kinase; (2) phosphotransacetylase; (3) carbon monoxide dehydrogenase; (4) methyltransferase; (5) methylene-THF reductase; (6) methylene-THF dehydrogenase; (7) methenyl-THF cyclohydrolase; (8) formyl-THF synthetase; (9) formate dehydrogenase. THF is tetrahydrofolate, Pi is PO4=, corrinoid/FeS is a corrinoid containing iron-sulfur protein, and [H] is reducing equivalents.

Methanogenesis: Syntrophic Metabolism

22

Schink, 2000). C. ultunense is also capable of both syntrophic acetate oxidation and acetate formation from H2 and CO2, but acetogenesis occurred only in cell suspensions (Schnu¨rer et al., 1997). The enzymes of the Wood–Ljungdahl pathway were detected in C. ultunense grown acetogenically and syntrophically except methylenetetrahydrofolate reductase, which was detectable acetogenically but not syntrophically (Schnu¨rer et al., 1997). T. lettingae does not contain genes for acetyl-CoA synthase/carbon monoxide dehydrogenase, which suggests that a pathway other than the Wood–Ljungdahl pathway is used (Hattori, 2008).

5.2

Propionate Metabolism

Two pathways for propionate metabolism are known, the methylmalonyl-CoA pathway and a novel dismutation pathway (> Fig. 2). The methylmalonyl-CoA pathway is found in many syntrophic propionate oxidizers including Syntrophobacter species (Boone and Bryant, 1980; Chen et al., 2005; Harmsen et al., 1998; Wallrabenstein et al., 1995), Desulfotomaculum thermobenzoicum subsp. thermosyntrophicum (Plugge et al., 2002), Pelotomaculum thermopropionicum (Imachi et al., 2002), and Pelotomaculum schinkii (de Bok et al., 2005). The dismutation pathway has been detected only in Smithella propionica (de Bok et al., 2001; Liu et al., 1999). S. propionica produces acetate and butyrate from propionate (de Bok et al., 2001). To explain the unusual labeling patterns observed in acetate and butyrate when different position-labeled propionate compounds were used, de Bok et al. (2001) concluded that two propionate molecules must condense to form a six-carbon intermediate, which is then rearranged to a 3-ketohexanoic acid intermediate before it is cleaved to form butyrate and acetate (> Fig. 2a). The enzymes involved in these novel reactions are not known. The methylmalonyl-CoA pathway (> Fig. 2b), also called the randomizing pathway, involves the activation of propionate to propionyl-CoA by transfer of a CoA group from acetyl-CoA and the synthesis of methylmalonyl-CoA by transfer of a carboxyl group from oxaloacetate by a transcarboxylase (Houwen et al., 1990). Methylmalonyl-CoA is then rearranged to form succinyl-CoA, which is oxidized via fumarate, oxaloacetate and pyruvate to acetate. The pathway predicts that one ATP is made by substrate-level phosphorylation per propionate degraded. Genomic and proteomic analyses show that the methylmalonyl-CoA pathway is operative in P. thermopropionicum (Kosaka et al., 2006). The production of hydrogen (or formate) from electrons derived from the oxidation of succinate is energetically difficult. Succinate reduced cytochrome b in membranes of Syntrophobacter fumaroxidans, and 2-(heptyl)-4-hydroxyquinoline-N-oxide inhibited succinate oxidation, suggesting the involvement of reverse electron transport (van Kuijk et al., 1998). Molar growth yields indicate that S. fumaroxidans synthesizes two-thirds of an ATP per fumarate when hydrogen is the electron donor. This observation suggests that S. fumaroxidans uses two-thirds of an ATP to drive hydrogen production from succinate when grown syntrophically with propionate; this leaves about one-third of an ATP available to support growth. The free energy change needed for irreversible ATP synthesis is estimated to be about
70 kJ mol
1 (Schink, 1997). If 3–5 protons are used to make ATP by the ATP synthase, then the minimum free energy change needed to form ATP in increments is
23 to
14 kJ mol
1 (Schink, 1997). This analysis predicts that syntrophic propionate metabolism should have a free energy change of about
20 kJ mol
1 to allow for the net synthesis of one-third of an ATP. Measured free energy changes during syntrophic propionate metabolism by S. fumaroxidans lower than
30 kJ mol
1 have been observed (Scholten and Conrad, 2000), which is in agreement with the energetic model.

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. Figure 2 Two pathways for syntrophic propionate metabolism. (a) The dismutation pathway for the metabolism of propionate by Smithella propionica. The carbons in each original propionate are labeled. The enzymes involved in this pathway have yet to be described and CoA esters of the compounds shown may be involved. This figure was adapted from de Bok et al. (2001). (b) The methylmalonyl-CoA pathway for propionate metabolism, found in P. thermopropionicum, was adapted from Kosaka et al. (2006). The enzymes involved are as follows: (1) propionate CoA transferase; (2) propionyl-CoA: oxaloacetate transcarboxylase; (3) methylmalonyl-CoA epimerase; (4) methylmalonyl-CoA mutase; (5) succinyl-CoA synthetase; (6) succinate dehydrogenase/fumarate reductase; (7) fumarate hydratase; (8) malate dehydrogenase; (9) pyruvate dehydrogenase; (10) pyruvate: formate lyase; (11) acetyl-CoA synthase; and (12) acetate kinase. Fd is ferredoxin and [H] is reducing equivalents.

Methanogenesis: Syntrophic Metabolism

22

However, under some growth conditions, the free energy available from syntrophic propionate metabolism <
10 kJ mol–1. Thus, we do not fully understand the bioenergetics of syntrophic propionate metabolism.

5.3

Butyrate Metabolism

Organisms capable of syntrophic butyrate metabolism include all species of Syntrophomonas (Lorowitz et al., 1989; McInerney et al., 1981; Sobieraj and Boone, 2006; Sousa et al., 2007; Wu et al., 2006a, b, 2007a, b; Zhang et al., 2004, 2005), Thermosyntropha lipolytica (Svetlitshnyi et al., 1996), and Syntrophothermus lipocalidus (Sekiguchi et al., 2000). Syntrophic butyrate metabolism proceeds via the b-oxidation pathway (> Fig. 3) (Wofford et al., 1986). Similar to syntrophic propionate metabolism, butyrate is activated to butyryl-CoA by the transfer of the

. Figure 3 The b-oxidation pathway for butyrate metabolism in Syntrophomonas wolfei, adapted from Wofford et al. (1986). The enzymes involved are: (1) CoA transferase; (2) acyl-CoA dehydrogenase; (3) enoyl-CoA hydratase; (4) L-(+)-3-hydroxybutyryl-CoA dehydrogenase; (5) 3-ketoacyl-CoA thiolase; (6) phosphotransacetylase; (7) acetate kinase. [H] is reducing equivalents.

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CoA group from acetyl-CoA; butyryl-CoA is then b-oxidized to two acetyl-CoA molecules (Wofford et al., 1986). One of the acetyl-CoA molecules is used to activate butyrate and the other is used for ATP synthesis. The oxidation of butyryl-CoA to crotonyl-CoA produces reduced electron transfer flavoprotein (E00 of
10 mV) (Sato et al., 1999) (> Fig. 3) and the oxidation of L-3-hydroxylbutyryl-CoA to 3-oxobutyryl-CoA produces NADH. Hydrogen production (E0 of
292 mV at 10 Pa H2) from electrons derived from NADH (E00 of
320 mV) (Thauer et al., 1977) is favorable at the partial pressures maintained by methanogens (about 1–10 Pa), but hydrogen production from the electrons from the oxidation of butyryl-CoA to crotonyl-CoA requires reverse electron transport (Wallrabenstein and Schink, 1994). About two-thirds of the ATP is needed to overcome this energy barrier, which leaves about one-third of an ATP available to support growth. The measured free energy changes available during syntrophic butyrate metabolism ranged from
5 to
17 kJ mol–1 (Dwyer et al., 1988; Jackson and McInerney, 2002), somewhat lower than that predicted to conserve energy (
14 to
23 kJ mol–1) (Schink, 1997).

5.4

Benzoate Metabolism

Syntrophic benzoate degraders include three species of Syntrophus: S. buswellii, S. gentianae and S. aciditrophicus, and Sporotomaculum syntrophicum, Pelotomaculum terephthalicicum and Pelotomaculum isophthalicicum (McInerney et al., 2008). The reduction of benzoyl-CoA represents a considerable energy barrier for anaerobic microorganisms because the midpoint potential of the first electron transfer is about
1.8 V (Boll and Fuchs, 1998; Boll et al., 2000; Heider and Fuchs, 1997a), which is well below that of most physiological electron donors (
0.4 V) (Boll and Fuchs, 1998). In Thauera aromatica, benzoyl-CoA reduction requires the hydrolysis of two ATP molecules per electron pair to overcome this barrier (Boll et al., 1997). Geobacter metallireducens (Wischgoll et al., 2005), Desulfococcus multivorans (Peters et al., 2004) and S. aciditrophicus (McInerney et al., 2007) most likely use a completely different type of enzyme to reduce benzoyl-CoA. Previous studies detected 2-hydroxycyclohexane carboxylate, cyclohex-1-ene carboxylate and pimelate in culture fluids of S. aciditrophicus grown with benzoate and the enzyme activities needed to convert cyclohex-1-ene carboxyl-CoA to pimelyl-CoA in cell-free extracts of S. aciditrophicus (> Fig. 4) (Elshahed et al., 2001). The intermediates and enzyme activities detected were consistent with the metabolism of cyclohex1-ene carboxyl-CoA to pimelyl-CoA by the pathway found in Rhodopseudomonas palustris (Harwood et al., 1998). However, genes homologous to those involved in benzoate metabolism in R. palustris were not detected in the S. aciditrophicus genome (McInerney et al., 2007). Interestingly, the genome of S. aciditrophicus contains genes with high homology to those in the T. aromatica genome (> Fig. 4) (McInerney et al., 2007). The genes for the cyclohex1,5-diene carboxyl-CoA hydratase and the 6-oxocyclohex-1-ene carboxyl-CoA hydrolase of S. aciditrophicus have been cloned and expressed in Escherichia coli (Kuntze et al., 2008; Peters et al., 2007). Enzymatic analysis showed that the S. aciditrophicus cyclohex-1,5-diene carboxylCoA hydratase converted cyclohex-1,5-diene carboxyl-CoA to 6-hydroxycyclohex-1-ene carboxyl-CoA, and that S. aciditrophicus 6-oxocyclohex-1-ene carboxyl-CoA hydrolase made 3-hydroxypimelyl-CoA from 6-oxocyclohex-1-ene carboxyl-CoA. Thus, it appears that S. aciditrophicus uses a two-electron reduction reaction to convert benzoyl-CoA to cyclohex1,5-diene carboxyl-CoA, which is then metabolized to 3-hydroxypimelyl-CoA in a manner analogous to that found in T. aromatica.

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. Figure 4 Pathway for syntrophic benzoate metabolism adapted from McInerney et al. (2007). The enzymes involved are: (1) benzoyl-CoA ligase; (2) benzoyl-CoA reductase; (3) cyclohex-1-carboxyl-CoA hydratase; (4) 2-hydroxycyclohexane carboxyl-CoA dehydrogenase; (5) 2-oxocyclohexanecarboxyl-CoA hydrolase; (6) pimelyl-CoA dehydrogenase; (7) enoyl-CoA hydratase; (8) cyclohex1,5-dienoyl-CoA hydratase; (9) 6-hydrocyclohex-1-ene-carboxyl-CoA dehydrogenase; (10) 6-oxocyclohex-1-ene-carboxyl-CoA hydrolase; (11) b-oxidation enzymes; (12) glutaconyl-CoA decarboxylase; (13) b-oxidation enzymes (see > Fig. 3 for more detail); (14) acetate kinase/ phosphotransacetylase. [H] are reducing equivalents. In step 2, the reduction of benzoyl-CoA is an ATP-dependent reaction in T. aromatica and R. palustris, while in S. aciditrophicus membrane potential rather than ATP hydrolysis may drive the electron transfer needed for ring reduction.

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It is not clear yet how net energy is conserved during syntrophic benzoate metabolism. Benzoate activation, benzoyl-CoA reduction and hydrogen production from acyl-CoA intermediates require more ATP (greater than 4 ATP per benzoate) than is produced by the known ATP-producing reactions, e.g., substrate-level phosphorylation from acetate (3 ATPs), proton translocation by a membrane-bound pyrophosphatase (equivalent to 1/3 ATP per benzoate) (Scho¨cke and Schink, 1998), and the decarboxylation of glutaconyl-CoA linked to sodium-ion translocation (1/3 of an ATP per benzoate) (Beatrix et al., 1990; Scho¨cke and Schink, 1998). Nevertheless, syntrophic benzoate degraders grow. The measured free energy changes during syntrophic benzoate metabolism range from about
30 to
45 kJ of energy (Scho¨cke and Schink, 1997; Warikoo et al., 1996), which suggest that about one-third of an ATP or more could be formed. In addition to the bioenergetics, there is still much that we do not understand of syntrophic benzoate metabolism. Cyclohexane carboxylate accumulates to very high concentration during syntrophic benzoate metabolism (Elshahed et al., 2001). Is it possible that another mechanism for benzoyl-CoA reduction exists in S. aciditrophicus, which may be energetically more favorable? Cyclohexane carboxylate and benzoate formation were observed when S. aciditrophicus was grown with crotonate (Mouttaki et al., 2007). Intermediates detected during crotonate metabolism were the same as those detected during syntrophic benzoate metabolism, suggesting that the pathway for benzoate metabolism is reversible and operates at high energy efficiency.

6

Potential Mechanisms for Reverse Electron Transport

Several mechanisms have been proposed for reverse electron transport during syntrophic metabolism (> Fig. 5). S. wolfei contains menaquinone (Wallrabenstein and Schink, 1994) which could function as the electron carrier between a membrane-associated acyl-CoA dehydrogenase and a cytoplasmically-oriented hydrogenase (Schink and Friedrich, 1994) (> Fig. 5a). In this model, inward movement of protons by the quinone loop is used to drive reverse electron transport. The second model (> Fig. 5b) directly links electron flow from the membrane-bound acyl-CoA dehydrogenase to an externally oriented hydrogenase (Schink and Friedrich, 1994). The consumption of protons on the outside of the cell membrane drives reverse electron transport. The external orientation of the hydrogenase in S. wolfei was confirmed by CuCl2 inhibition studies (Wallrabenstein and Schink, 1994). A third variation is possible if a soluble acyl-CoA dehydrogenase is used (> Fig. 5c). In this model, a membrane-bound oxidoreductase would be necessary to aid electron transfer to the hydrogenase. This model could employ menaquinone or another electron carrier and would involve the proton gradient as the driving force for reverse electron transport. Another possibility to produce H2 from thermodynamically difficult substrates was recently proposed by Prof. Buckel’s laboratory and confirmed by Prof. Thauer’s laboratory in Clostridium kluyveri (Herrmann et al., 2008; Li et al., 2008). C. kluyveri ferments ethanol and acetate to butyrate and small amounts of H2. A soluble enzyme complex in C. kluyveri couples the energetically favorable reduction of crotonyl-CoA to butyryl-CoA by NADH with the unfavorable reduction of ferredoxin (Fd) by NADH (> Fig. 5d) (1): Crotonyl-CoA + Fdox + 2NADH + 2H+! Butyryl-CoA + Fdred + 2NAD+ (1) This enzyme complex was purified and shown to be FAD dependent (Li et al., 2008). Once FADH2 is formed, there is a bifurcation of electron flow with some electrons used for the

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. Figure 5 Proposed models for reversed electron transport chains. (a) and (b) adapted from Schink and Friedrich (1994), (d) adapted from Li et al. (2008), and (e) adapted from Schirawski and Unden (1998). DH, dehydrogenase; Hase, hydrogenase; MK, menaquinone; Fd is ferredoxin; ?, unknown membrane bound oxidoreductase; ox:red, oxidoreductase; in, cytoplasm; out, outside cytoplasmic membrane.

exergonic reduction of crotonyl-CoA (E 00 =
10 mV), which drives the endergonic reduction of ferredoxin (E 00 =
410 mV). The reversal of this reaction could accomplish reverse electron transport during syntrophic oxidation. A possible model for reverse electron transport during syntrophic propionate metabolism is suggested by studies in Bacillus subtilis (> Fig. 5e) (Schirawski and Unden, 1998). B. subtilis has menaquinone (E 00 of
70 mV) and not ubiquinone (E 00 of + 100 mV) so the oxidation of succinate to fumarate (E 00 of + 33 mV) is coupled to menaquinone reduction and requires reverse electron transport. The binding site for menaquinone of the membrane-bound cytochrome b was located close to the outside of the cell membrane, which would allow the inward movement of protons if menaquinone is oxidized on the cytoplasmic side of the membrane. In syntrophic propionate metabolism, menaquinone oxidation could be linked to a membranebound hydrogenase or formate dehydrogenase. S. fumaroxidans has a membrane-bound succinate dehydrogenase (van Kuijk et al., 1998) and formate dehydrogenases and hydrogenases that are membrane-associated (de Bok et al., 2002a, b, 2003). These observations are consistent with the a reverse electron transport system as found in Bacillus subtilis (Schirawski and Unden, 1998).

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Research Needs

The concept of a minimum free energy change for energy conservation provides the framework to understand how bacteria exploit small free energy changes. Pathways for syntrophic metabolism of fatty acids predict that ATP can be synthesized at increments of about one-third of an ATP, which is consistent with the measured free energy changes observed for the syntrophic metabolism of these compounds. However, there is still much that we do not understand. For instance, how does net ATP synthesis occur during syntrophic benzoate metabolism and why, in some cases, are free energy changes less than the predicted minimum free energy change needed for energy conservation? We know very little about the biochemistry of reverse electron transport. Genome sequences of several organisms capable of syntrophic metabolism are available and analysis of these sequences may provide insights into the bioenergetics of syntrophy. Genomic analyses suggest that organisms capable of syntrophic metabolism have multiple mechanisms to generate ion gradients and use different mechanisms for reverse electron transport. Further work on the role of these membrane complexes in syntrophic metabolism is needed. More research is needed to understand how bacteria exploit small free energy changes for growth. Also unresolved is how reversible metabolic pathways found in some syntrophic metabolizers are linked to ATP synthesis in each direction. Many syntrophic associations are highly organized, multicellular structures with the partners in close physical proximity to each other. We know very little about the molecular mechanisms involved in the formation and maintenance of these catalytic units. Regulatory mechanisms that control the development of attached consortia most likely are similar to those involved in biofilm formation. We are beginning to unravel the molecular and biochemical details involved in the syntrophic lifestyle. The combination of computational with functional genomic approaches may allow us to interrogate the regulatory mechanisms involved in establishing and maintaining multispecies associations in order to quantify and predict the behavior of microorganisms and microbial communities in natural ecosystems. A thorough understanding of the formation and structure of dense microbial aggregates is essential for application of methanogenesis. Only in bioreactors in which methanogenic communities operate in dense aggregates can anaerobic wastewater treatment take place at a high volumetric rate. Many syntrophic associations still need to be discovered. Besides syntrophic oxidations of the compounds discussed here, syntrophic interactions may also play an important role in the degradation of compounds that are considered to be easily fermentable, e.g., sugars as shown by Krumholz and Bryant (1986) and Mu¨ller et al. (2008). Besides freshwater environments, sulfate-depleted marine sediments are also important methanogenic environments (Colwell et al., 2008). Syntrophic interactions in these marine methanogenic environments have not been studied yet. Only recently, propionate and butyrate-degrading methanogenic communities from marine samples have been described (Kendall et al., 2006). In order to identify still unknown syntrophic interactions, metagenomic approaches may be of great help, especially if key enzymes and the corresponding genes, e.g., the ones involved in reversed electron transfers, have been studied in depth.

Acknowledgments The work of M. J. McInerney and R. P. Gunsalus was supported by contracts DE-FG0296ER20214 and DE-FG02-08ER64689 from the U.S. Department of Energy.

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metabolism of 4-hydroxybenzoate by Rhodopseudomonas palustris. J Bacteriol 171(1): 1–7. Mountfort DO, Bryant MP (1982) Isolation and characterization of an anaerobic syntrophic benzoatedegrading bacterium from sewage sludge. Arch Microbiol 133(4): 249–256. Mouttaki H, Nanny MA, McInerney MJ (2007) Cyclohexane carboxylate and benzoate formation from crotonate in Syntrophus aciditrophicus. Appl Environ Microbiol 73(3): 930–938. Mu¨ller N, Griffin BM, Stingl U, Schink B (2008) Dominant sugar utilizers in sediment of Lake Constance depend on syntrophic cooperation with methanogenic partner organisms. Environ Microbiol 10: 1501–1511. Pavlostathis SG, Giraldo-Gomez E (1991) Kinetics of anaerobic treatment: a critical review. Crit Rev Environ Control 21(5): 411–490. Peters F, Rother M, Boll M (2004) Selenocysteinecontaining proteins in anaerobic benzoate metabolism of Desulfococcus multivorans. J Bacteriol 186(7): 2156–2163. Peters F, Shinoda Y, McInerney MJ, Boll M (2007) Cyclohexa-1,5-diene-1-carbonyl-coenzyme A (CoA) hydratases of Geobacter metallireducens and Syntrophus aciditrophicus: evidence for a common benzoyl-CoA degradation pathway in facultative and strict anaerobes. J Bacteriol 189(3): 1055–1060. Plugge CM, Balk M, Stams AJ (2002) Desulfotomaculum thermobenzoicum subsp. thermosyntrophicum subsp. nov., a thermophilic, syntrophic, propionateoxidizing, spore-forming bacterium. Int J Syst Evol Microbiol 52(2): 391–399. Reguera G, McCarthy KD, Mehta T, Nicoll JS, Tuominen MT, Lovley DR (2005) Extracellular electron transfer via microbial nanowires. Nature 435(7045): 1098–1101. Sato K, Nishina Y, Setoyama C, Miura R, Shiga K (1999) Unusually high standard redox potential of acrylylCoA/propionyl-CoA couple among enoyl-CoA/ acyl-CoA couples: a reason for the distinct metabolic pathway of propionyl-CoA from longer acylCoAs. J Biochem 126(4): 668–675. Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61(2): 262–280. Schink B, Friedrich M (1994) Energetics of syntrophic fatty acid oxidation. FEMS Microbiol Rev 15(2–3): 85–94. Schink B, Philipp B, Mu¨ller J (2000) Anaerobic degradation of phenolic compounds. Naturwissenschaften 87(1): 12–23. Schink B, Stams AJM (2006) Syntrophism among prokaryotes. In The Prokaryotes: An Evolving Electronic Resource for the Microbiological Community.

M Dworkin, S Falkow, E Rosenberg, KH Schleifer and E Stackebrandt (eds.). 3rd edn. Vol. 2. New York: Springer, pp. 309–335. Schirawski J, Unden G (1998) Menaquinone-dependent succinate dehydrogenase of bacteria catalyzes reversed electron transport driven by the proton potential. Eur J Biochem 257(1): 210–215. Schnu¨rer A, Svensson BH, Schink B (1997) Enzyme activities in and energetics of acetate metabolism by the mesophilic syntrophically acetate-oxidizing anaerobe Clostridium ultunense. FEMS Microbiol Lett 154(2): 331–336. Scho¨cke L, Schink B (1997) Energetics of methanogenic benzoate degradation by Syntrophus gentianae in syntrophic coculture. Microbiology 143: 2345–2351. Scho¨cke L, Schink B (1998) Membrane-bound protontranslocating pyrophosphatase of Syntrophus gentianae, a syntrophically benzoate-degrading fermenting bacterium. Eur J Biochem 256(3): 589–594. Scholten JC, Conrad R (2000) Energetics of syntrophic propionate oxidation in defined batch and chemostat cocultures. Appl Environ Microbiol 66(7): 2934–2942. Sekiguchi Y, Kamagata Y, Nakamura K, Ohashi A, Harada H (2000) Syntrophothermus lipocalidus gen. nov., sp. nov., a novel thermophilic, syntrophic, fatty-acid-oxidizing anaerobe which utilizes isobutyrate. Int J Syst Evol Microbiol 50(2): 771–779. Sieber J, Gunsalus RP, Rohlin L, McInerney MJ, Sims DR, Han C, Kim E, Lykidis A, Lapidus AL (2008) Genomic insights into syntrophic fatty acid metabolism: electron transfer processes of Syntrophomonas wolfei. American Society of Microbiology 108th General Meeting. Boston, MA, Abst. I-002, p. 071. Sobieraj M, Boone DR (2006) Syntrophomonadaceae. In The Prokaryotes: An Evolving Electronic Resource for the Microbiological Community, M Dworkin, S Falkow, E Rosenberg, KH Schleifer, and E Stackebrandt (eds.). 3rd edn, vol. 4., New York: Springer, pp. 1041–1046. Sousa DZ, Smidt H, Alves MM, Stams AJ (2007) Syntrophomonas zehnderi sp. nov., an anaerobe that degrades long-chain fatty acids in co-culture with Methanobacterium formicicum. Int J Syst Evol Microbiol 57(3): 609–615. Svetlitshnyi V, Rainey F, Wiegel J (1996) Thermosyntropha lipolytica gen. nov., sp. nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing short- and long-chain fatty acids in syntrophic coculture with a methanogenic archaeum. Int J Syst Bacteriol 46(4): 1131–1137. Szewzyk U, Schink B (1989) Degradation of hydroquinone, gentisate, and benzoate by a fermenting bacterium in pure or defined mixed culture. Arch Microbiol 151(6): 541–545.

Methanogenesis: Syntrophic Metabolism Thauer RK, Jungermann K, Decker K (1977) Energy conservation in chemotrophic anaerobic bacteria. Bacteriol Rev 41(1): 100–180. Thiele JH, Zeikus JG (1988) Control of interspecies electron flow during anaerobic digestion: significance of formate transfer versus hydrogen transfer during syntrophic methanogenesis in flocs. Appl Environ Microbiol 54(1): 20–29. van Kuijk BL, Schlosser E, Stams AJ (1998) Investigation of the fumarate metabolism of the syntrophic propionate-oxidizing bacterium strain MPOB. Arch Microbiol 169(4): 346–352. Wallrabenstein C, Hauschild E, Schink B (1995) Syntrophobacter pfennigii sp. nov., new syntrophically propionate-oxidizing anaerobe growing in pure culture with propionate and sulfate. Arch Microbiol 164(5): 346–352. Wallrabenstein C, Schink B (1994) Evidence of reversed electron transport in syntrophic butyrate or benzoate oxidation by Syntrophomonas wolfei and Syntrophus buswellii. Arch Microbiol 162(1): 136–142. Warikoo V, McInerney MJ, Robinson JA, Suflita JM (1996) Interspecies acetate transfer influences the extent of anaerobic benzoate degradation by syntrophic consortia. Appl Environ Microbiol 62(1): 26–32. Wischgoll S, Heintz D, Peters F, Erxleben A, Sarnighausen E, Reski R, Van Dorsselaer A, Boll M (2005) Gene clusters involved in anaerobic benzoate degradation of Geobacter metallireducens. Mol Microbiol 58(5): 1238–1252. Wofford NQ, Beaty PS, McInerney MJ (1986) Preparation of cell-free extracts and the enzymes involved in fatty acid metabolism in Syntrophomonas wolfei. J Bacteriol 167(1): 179–185. Wu C, Dong X, Liu X (2007a) Syntrophomonas wolfei subsp. methylbutyratica subsp. nov., and assignment of Syntrophomonas wolfei subsp. saponavida to

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Syntrophomonas saponavida sp. nov. comb. nov. Syst Appl Microbiol 30(5): 376–380. Wu C, Dong X, Liu X (2007b) Syntrophomonas wolfei subsp. methylbutyratica subsp nov., and assignment of Syntrophomonas wolfei subsp. saponavida to Syntrophomonas saponavida sp. nov. comb. nov. Syst Appl Microbiol. doi:10.1016/j.syapm.2006.12.001. Wu C, Liu X, Dong X (2006a) Syntrophomonas erecta subsp. sporosyntropha subsp. nov., a spore-forming bacterium that degrades short chain fatty acids in co-culture with methanogens. Syst Appl Microbiol 29(6): 457–462. Wu C, Liu X, Dong X (2006b) Syntrophomonas cellicola sp. nov., a spore-forming syntrophic bacterium isolated from a distilled-spirit-fermenting cellar, and assignment of Syntrophospora bryantii to Syntrophomonas bryantii comb. nov. Int J Syst Evol Microbiol 56(10): 2331–2335. Zhang C, Liu X, Dong X (2004) Syntrophomonas curvata sp. nov., an anaerobe that degrades fatty acids in coculture with methanogens. Int J Syst Evol Microbiol 54(3): 969–973. Zhang C, Liu X, Dong X (2005) Syntrophomonas erecta sp. nov., a novel anaerobe that syntrophically degrades short-chain fatty acids. Int J Syst Evol Microbiol 55(2): 799–803. Zhilina TN, Zavarzina DG, Kolganova TV, Turova TP, Zavarzin GA (2005) ‘‘Candidatus Contubernalis alkalaceticum,’’ an obligately syntrophic alkaliphilic bacterium capable of anaerobic acetate oxidation in a coculture with Desulfonatronum cooperativum. Microbiology 74(6): 800–809. Zindel U, Freudenberg W, Rieth M, Andreesen JR, Schnell J, Widdel F (1988) Eubacterium acidaminophilum sp. nov., a versatile amino acid-degrading anaerobe producing or utilizing H2 or formate. Arch Microbiol 150(3): 254–266.

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23 Biochemistry of Acetotrophic Methanogenesis J. G. Ferry Department of Biochemistry and Molecular Biology, The Pennsylvania State University, PA, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 358 2 Coenzymes and Cofactors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 359 3 Synthesis of CH3-H4SPT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 360 4 Conversion of CH3-H4SPT to Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 5 Energy Conservation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 364 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 366

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Abstract: Two-thirds of the estimated one billion metric tones of methane produced each year in the Earth’s biosphere derives from the methyl group of acetate via the acetotrophic pathway. The pathway is best understood in species from the genus Methanosarcina, one of only two genera known to obtain energy for growth by converting acetate to methane and carbon dioxide. The pathway commences by activation of acetate to acetyl-CoA of which the C–C and C–S bonds are cleaved yielding methyl and carbonyl groups. The methyl group is ultimately transferred to coenzyme M that is reductively demethylated to methane with electrons derived from oxidation of the carbonyl group of acetate. Cells obtain energy for growth by the transfer of electrons through a membrane-bound electron transport chain coupled to generation of an ion gradient that drives ATP synthesis. This review summarizes the current biochemical understanding of the pathway contrasting marine and freshwater Methanosarcina species with a focus on the mechanism of key enzymes.

1

Introduction

Methane is the end product of the decomposition of complex organic matter in diverse O2-free (anaerobic) environments, producing nearly 1 billion metric tons of methane each year. The process is an essential link in Earth’s carbon cycle (> Fig. 1). In the cycle, CO2 is fixed into complex organic matter by photosynthesis (step 1) and in aerobic zones is oxidized back to CO2 by O2-requiring microbes (step 2). A portion of the organic matter is deposited in a variety of anaerobic environments where diverse anaerobes decompose the organic matter (step 3) to products that are substrates for methane-producing species (steps 4 and 5). Some of the methane is utilized by sulfate- or nitrate-reducing anaerobic methane oxidizers (step 6) (Thauer and Seigo, 2007) and the remainder escapes into aerobic habitats where it is oxidized to CO2 by O2-requiring methylotrophic microbes (step 7). Approximately one-third of the methane produced in Earth’s biosphere is generated by the reduction of CO2 with electrons derived from the oxidation of H2 (> Fig. 1, step 4): CO2 þ 4H2 ! CH4 þ 2H2

ð1Þ

The remaining two-thirds originates from the methyl group of acetate (> Fig. 1, step 5) by the ‘‘aceticlastic’’ pathway: CH3 COO þ Hþ ! CH4 þ CO2

. Figure 1 The global carbon cycle.

ð2Þ

Biochemistry of Acetotrophic Methanogenesis

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Only two genera of methanogens (> Fig. 2), Methanosarcina (Msr.) and Methanosaeta (Mse.), contain acetate-utilizing species (Ferry and Kastead, 2007). Methanosaeta (previously Methanothrix) species have a higher affinity for acetate than Methanosarcina and dominate in methanogenic habitats with low concentrations of acetate. The biochemistry of the aceticlastic pathway has been investigated primarily in the freshwater Methanosarcina species Msr. barkeri, Msr. mazei and Msr. thermophila and the marine isolate Msr. acetivorans.

2

Coenzymes and Cofactors

> Figure

2 illustrates the structures of cofactors and coenzymes involved in the aceticlastic pathway. Factor III, tetrahydromethanopterin (H4MPT), coenzyme M (HS-CoM) and factor F430 (F430) all are involved in transfer of the methyl group of acetate to methane. Factor III binds the methyl group as an upper axial ligand of the cobalt atom analogous to vitamin B12 and has a similar structure except the lower axial ligand is a 5-hydroxybenzimidazolyl

. Figure 2 Cofactors and coenzymes utilized in the aceticlastic pathway of methanogenesis.

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base. H4MPT carries a methyl group bound to the 5N position. Methanosarcina species synthesize tetrahydrosarcinapterin (H4SPT) that has a structure similar to H4MPT except for an added terminal a-linked glutamate. Coenzyme M (HS-CoM) carries a methyl group attached to the sulfur atom. F430 is a nickel-containing corphinoid tetrapyrrole that binds a methyl group to the metal atom. Coenzyme B (CoB-SH) and methanophenazine (MP) are electron carriers.

3

Synthesis of CH3-H4SPT

Biochemical and bioinformatic evidence indicates that the core methyl transfer steps leading from the methyl group of acetate to methane are similar in all freshwater and marine species (> Fig. 3). Reactions involved in transfer of the methyl group from acetate to produce CH3– H4SPT (3, 4) are catalyzed by enzymes with homologs widely dispersed in the Bacteria domain. CH3 COO þ ATP ! CH3 CO2 PO2 3 þ ADP

ð3Þ

CH3 CO2 PO2 3 þ HS-CoA ! CH3 COSCoA þ Pi

ð4Þ

CH3 COSCoA þ H4 SPT þ H2 O þ Fdo ! CH3H4 SPT þ Fdr þ CO2 þ HSCoA

ð5Þ

The structure and function of enzymes catalyzing these reactions have been investigated in significant detail, the understanding of which has impacted the broader field of prokaryotic biology in view of the fact that paralogs are wide-spread in diverse anaerobes from the Bacteria domain. Acetate kinase (> Fig. 3, Ack) and phosphotransacetylase (> Fig. 3, Pta) catalyze reactions (3, 4) that together activate acetate to acetyl-CoA which is the substrate for CO dehydrogenase/ acetyl-CoA (> Fig. 3, Cdh), the central enzyme in the pathway catalyzing the reaction shown in Equation 5. Overproduction in Escherichia coli of acetate kinase and phosphotransacetylase from Msr. thermophila has established these enzymes as models for investigation. The crystal structure (Buss et al., 2001) of the acetate kinase suggests that it is the founding member of the ASKHA (Acetate and Sugar Kinase/Hsc70/Actin) superfamily of phosphotransferases. Prior to the structure of the Msr. thermophila enzyme, the first for any acetate kinase, a triple displacement mechanism involving two covalent phosphoenzyme intermediates was proposed for the acetate kinase from E. coli (Spector, 1980). However, kinetic and biochemical studies of the wild-type versus site-specific amino acid variants of the Msr. thermophila enzyme support a direct in-line mechanism for transfer of the phosphate from ATP to acetate (Gorrell and Ferry, 2007). The crystal structure of the Msr. thermophila phosphotransacetylase with CoA-SH bound has identified the active site residues leading to a proposal for the catalytic mechanism (Lawrence et al., 2006). In the mechanism, base catalysis generates -S-CoA followed by nucleophilic attack of the thiolate anion on the carbonyl carbon of acetyl phosphate yielding acetyl-CoA and inorganic phosphate. In Methanosaeta species, acetate is converted to acetyl-CoA in one reaction (6) catalyzed by acetyl coenzyme A synthetase (Eggen et al., 1991). CH3 COO þ ATP þ CoASH ! CH3 COSCoA þ AMP þ PiPi

ð6Þ

The CO dehydrogenase acetyl-CoA Synthase (Cdh) cleaves the C–C and C–S bonds of acetyl-CoA (5) yielding methyl and carbonyl groups (Ferry, 1995). The methyl group is

Biochemistry of Acetotrophic Methanogenesis

. Figure 3 (Continued)

23

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transferred to H4SPT for eventual conversion to methane and the carbonyl group is oxidized to CO2 with the electrons transferred to a 2x[4Fe-4S] ferredoxin. In addition to aceticlastic methanogens, diverse anaerobes from the Bacteria domain utilize Cdh in energy-yielding pathways generating acetyl-CoA with CoA-SH, a methyl group and CO2 plus a pair of electrons. The acetyl-CoA is further metabolized to acetate catalyzed by phosphotransacetylase and acetate kinase producing ATP. The Cdh from methanogens has been biochemically characterized from both Methanosarcina and Methanosaeta species. Most structure and function studies have been performed with the enzymes from Msr. thermophila and Msr. barkeri. In both species the enzyme is a complex comprised of five-subunits (a,b,g,d,e) resolvable by detergent treatment into a Ni/Fe-S component (a and e subunits), a Co/Fe-S component (g and d subunits), and the b subunit. The resolved Ni/Fe-S component catalyzes the reversible oxidation of CO to CO2 utilizing a 2x[4Fe-4S] ferredoxin as the redox partner consistent with a role for this component in oxidizing the carbonyl group of acetyl-CoA and reducing ferredoxin. A crystal structure for the Ni/Fe-S component from Msr. barkeri (Gong et al., 2008) reveals the a subunit containing a pseudocubane Ni-3Fe-4S cluster of which an acid-labile sulfide is bridged to an exogenous Fe atom. Electron density adjacent to this structure is consistent with ligation of the carbon of CO to Ni and the oxygen hydrogen bonded to a water molecule that is a ligand of the exogenous Fe. Thus, the Ni and exogenous Fe is thought to play an activation role in breaking the C–O bond. Structural analysis of the e subunit revealed a potential FAD binding site consistent with the previously reported ability of the Ni/Fe-S component from Msr. barkeri to reduce FAD. Furthermore, the potential FAD binding site is located adjacent to a 4Fe-4S cluster in the a subunit. Thus, a function for the e subunit in transfer of electrons from the 4Fe-4S cluster to FAD is plausible. Another structural feature distinct from homologs of the Bacteria domain is the presence of two additional 4Fe-4S clusters in the a subunit that are postulated to function in transfer of electrons to ferredoxin. The b component of the Cdh of Msr. thermophila contains an ‘‘A’’ cluster that is the proposed site of acetyl-CoA cleavage or synthesis (Gencic and Grahame, 2003). A crystal structure has not been reported for the b component of Cdh; however, the structure of the homolog from Moorella thermoacetica (Ragsdale, 2004), a non-methanogen from the Bacteria domain, reveals the ‘‘A’’ cluster with a 4Fe-4S center bridged to a Ni via sulfur atoms and an undisclosed metal proximal to the 4Fe-4S center. The identity of the proximal metal is equivocal, although Ni is a likely candidate. Assembly of acetyl-CoA is proposed to occur on the Ni distal to the 4Fe-4S center. Ni and Fe K fluorescence XANES and EXAFS analyses of the b component from Msr. barkeri, expressed as a C-terminally shortened form, identified an 4Fe-4S cluster and a

. Figure 3 Proposed acetotrophic pathways of methanogenesis by the freshwater species Msr. mazei (panel A) and the marine isolate Msr. acetivorans (panel B). Ack, acetate kinase; Pta, phosphotransacetylase; CoA-SH, coenzyme A; H4SPT, tetrahydrosarcinapterin; Fdr, reduced ferredoxin; Fdo, oxidized ferredoxin; Cdh, CO dehydrogenase/acetyl-CoA synthase; CoM-SH, coenzyme M; Mtr, CH3-H4SPT:CoM-SH methyltransferase; CoB-SH, coenzyme B; Cam, carbonic anhydrase; Ech, H2-evolving hydrogenase; Vho, H2-consuming hydrogenase; Ma-Rnf, Msr. acetivorans Rnf; MP, methanophenazine; Hdr-DE, heterodisulfide reductase; Mrp, multiple resistance/pH regulation Na+/H+antiporter; Atp, H+-translocating A1A0 ATP synthase. Adapted from (Li et al., 2006).

Biochemistry of Acetotrophic Methanogenesis

23

ratio of 2 Ni per cluster (Gu et al., 2003). The results are consistent with an active-site structure containing a binuclear Ni-Ni bridged to the 4Fe-4S cluster by a cysteine thiolate similar to that proposed for the enzyme from M. thermoacetica. Redox titration experiments indicated that two electrons are required for activation of the enzyme in the process of forming an enzymeacetyl intermediate (Gencic and Grahame, 2008). The Co/Fe-S component of the Cdh from Methanosarcina species contains Factor III and is involved in transfer of the methyl group of acetyl-CoA to H4SPT (Ferry, 1995). The cobalt and Fe-S centers of the Co/Fe-S component from Msr. thermophila were investigated using EPR spectroscopy and spectroelectrochemistry showing that Factor III is maintained in the base-off state with a formal equilibrium reduction potential at pH 7.8 of 486 mV for the Co2+/Co1+couple that facilitates reduction of the Co2+ state by approximately 12 kcal/mol relative to base-on. Maintenance of the Co1+state is important since only Co1+can be methylated; thus, increasing the potential into this higher range is important since reduction of Co+2 with electrons diverted from the Ni/Fe-S component would otherwise be exergonic. The Co/Fe-S component also contains a [4Fe-4S] cluster with an midpoint potential at pH 7.8 of 502 mV, which is nearly isopotential with the Co2+/ Co1+couple and likely serves as the direct electron donor. The cdhD and cdhE genes which encode the d and g subunits have been cloned and sequenced (Maupin-Furlow and Ferry, 1996). The CdhE sequence contains a four-cysteine motif with the potential to bind a 4Fe-4S cluster. Further, CdhE overproduced in E. coli contains the cluster and a corrinoid cofactor with the benzimidazole base in the base-off configuration. The results are consistent with a role for the g subunit in transfer of the methyl group to H4SPT.

4

Conversion of CH3-H4SPT to Methane

The conversion of CH3–H4SPT to methane is common to all methanogenic pathways and requires three reactions catalyzed by a membrane-bound CH3–H4SPT: coenzyme M methyltransferase (> Eq. 7), CH3–CoM methylreductase (> Eq. 8) and heterodisulfide reductase (> Eq. 9). CH3THMPT þ HSCoM ! CH3SCoM þ THMPT

ð7Þ

CH3SCoM þ HSCoB ! CoMSSCoB þ CH4

ð8Þ

CoMSSCoB þ 2e þ 2Hþ ! HSCoB þ HSCoM

ð9Þ

The methyltransferase (> Fig. 3, Mtr) has been characterized from acetate-grown cells of Msr. mazei and other methanoarchaea that has been reviewed (Gottschalk and Thauer, 2001). The enzyme, comprised of eight non-identical subunits couples the exergonic methyl transfer to generation of a sodium ion gradient (high outside the cytoplasmic membrane) postulated to drive various energy-requiring reactions. The MtrA subunit containing Factor III is proposed to extend into the cytoplasm where the methyl group of CH3–H4SPT is transferred to the cobalt of Factor III catalyzed by MtrH. MtrE is postulated to demethylate Factor III of MtrA resulting in loss of the lower axial histidine ligand and inducing a conformational change transduced to MtrE that drives the translocation of sodium. The reaction shown in > Eq. 8 is catalyzed by methyl-CoM methylreductase (> Fig. 3, Mcr) that contains nickel in coenzyme F430. HS–CoB is the electron donor that when oxidized forms a disulfide bond with HS–CoM (CoM–S–S–CoB) in addition to methane. The only

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crystal structure available for the methylreductase is that from the CO2-reducing species Methanothermobacter marburgensis (Methanobacterium thermoautotrophicum strain Marburg) that shows the three non-identical subunits in a a2b2g2 arrangement (Ermler et al., 1997). A concerted mechanism is proposed in which CH3–S–CoM binds adjacent to the nickel atom of F430 before entry of HS-CoB in the narrow active site channel. Nucleophilic attack of Ni(I) on CH3–S–CoM forms a Ni(III)–CH3 intermediate and HS–CoM. In the next step, electrons are transferred from HS-CoM to Ni(III) producing the thiyl radical S–CoM and Ni(II)–CH3. The S–CoM is coupled with S–CoB to form CoB-S-S-CoM followed by one-electron reduction of Ni(II) to Ni(I) regenerating the active Ni(I) form of F430. In the final step, protonolysis of Ni(II)–CH3 in F430 produces methane. A two-subunit heterodisulfide reductase (> Fig. 3, Hdr) identified in acetate-grown cells of Msr. thermophila (Murakami et al., 2001) and Msr. barkeri (Hedderich et al., 2005) is consistent with a role for catalysis of the disulfide bond of CoM–S–S–CoB and regenerating the active sulfhydryl forms of the coenzymes (9). A methanophenazine analog, 2-hydroxyphenazine (Murakami et al., 2001), is an electron donor to the Msr. thermophila heterodisulfide reductase consistent with a role for methanophenazine in electron transport and generation of a proton gradient (> Fig. 3). The HdrE subunit of the Msr. barkeri enzyme contains cytochrome b that transfers electrons to a 4Fe-4S cluster in the catalytic subunit HdrD where reduction of CoM–S–S–CoB is proposed to take place in two one-electron steps (Hedderich et al., 2005). Involvement of a 4Fe-4S cluster is also proposed for the HdrDE from acetate-grown Msr. thermophila, although kinetic studies and inhibition experiments indicate that the order of electron flow is: methanophenazine ! high potential 4Fe-4S cluster ! low potential heme ! CoM–S–S–CoB (Murakami et al., 2001).

5

Energy Conservation

The heterodisulfide CoM–S–S–CoB is the terminal electron acceptor of a membrane-bound electron transport chain coupled to formation of an electrochemical ion gradient driving ATP synthesis. The ‘‘archaeal’’ A1A0-type ATP synthase is abundant in acetate-grown Msr. acetivorans (Li et al., 2007) and Msr. mazei (Hovey et al., 2005) consistent with a role in ATP synthesis. Although sequence analysis identifies a putative Na+binding site in the A1A0-type of methanogens, including Msr. acetivorans and Msr. mazei, the coupling ion appears to be H+(Pisa et al., 2007). A membrane-bound reduced ferredoxin (Fdr):CoM–S–S–CoB oxidoreductase system generates a electrochemical gradient which drives ATP synthesis in freshwater Methanosarcina species (> Fig. 3a) (Deppenmeier, 2004). The ferredoxin is reduced with electrons derived from oxidation of the carbonyl group of acetyl-CoA catalyzed by the Cdh. In the freshwater species Msr. barkeri, it is proposed that ferredoxin donates electrons to Ech hydrogenase which produces H2 and generates a proton gradient (high outside) supported by gene knockout experiments showing that Ech is essential for growth with acetate (Meuer et al., 2002). The proposed role of H2 as an electron transport intermediate is further supported by the presence of a cytochrome b-containing H2:heterodisulfide oxidoreductase complex characterized from acetate-grown Msr. barkeri. An uptake hydrogenase (Vho/Vht) is proposed to reoxidize H2 with transfer of electrons to methanophenazine that is the electron donor to the heterodisulfide reductase (Meuer et al., 2002) (> Fig. 3a). The heterodisulfide reductase purified from acetate-grown Msr. thermophila is reported to have hydrogenase activity (Simianu et al., 1998)

Biochemistry of Acetotrophic Methanogenesis

23

consistent with a role for H2 in the electron transport chain of this freshwater species. A role for Isf (iron-sulfur flavoprotein) in the transfer of electrons from ferredoxin to the membranebound electron transport chain has been proposed for Msr. thermophila (Latimer et al., 1996). In the freshwater species Msr. mazei the expression of genes encoding Ech are up regulated in acetate versus methanol-grown cells (Hovey et al., 2005) further supporting a role for this hydrogenase in acetate-utilizing freshwater Methanosarcina species. The role of an Isf homolog in the transfer of electrons from Ech to CoM–S–S–CoB in Msr. mazei has been postulated based on up regulation of the encoding gene in response to growth on acetate (Hovey et al., 2005). Msr. acetivorans, a marine species, has evolved a mechanism for oxidizing ferredoxin and reducing CoM–S–S–CoB that does not involve H2 (> Fig. 3b). A functional Ech hydrogenase is not encoded in the genome (Galagan et al., 2002) and acetate-grown cells have relatively little H2-dependent methylreductase activity (Nelson and Ferry, 1984). Based on quantitative proteomic and biochemical analyses, it is proposed that a homolog of Rnf (> Fig. 3b, Ma-Rnf) oxidizes ferredoxin in place of Ech and that methanophenazine mediates electron transfer between reduced Ma-Rnf and the heterodisulfide reductase HdrDE (Li et al., 2006). Rnf was first discovered in Rhodobacter capsulatus and shown to be a six-subunit membrane-bound electron transfer complex with homologs wide spread in the Bacteria domain that are proposed to couple electron transport to the generation of a Na+gradient (Boiangiu et al., 2005). The six subunits of the Msr. acetivorans Ma-Rnf complex are encoded in a transcriptional unit with two additional flanking ORF’s, one of which encodes a cytochrome c (Li et al., 2006). It is proposed that the Ma-Rnf complex and cytochrome c function to transport electrons from ferredoxin to methanophenazine coupled to generation of a sodium gradient (high outside the membrane). Subunits of a seven-subunit Na+/H+antiporter (> Fig. 3b, Mrp) are at least 30-fold more abundant in acetate versus methanol-grown Msr. acetivorans (Li et al., 2006) suggesting a role for this complex to exchange the Na+gradient for a proton gradient that drives ATP synthesis by the proton-dependent A1A0 ATP synthase (> Fig. 3b). The genomic sequence of Mse. thermophila indicates that core reactions converting the methyl group of acetate to methane are similar to that of freshwater and marine Methanosarcina species (Smith and Ingram-Smith, 2007). Conversely, genes encoding Ech and Rnf are absent in the genome of Mse. thermophila suggesting an alternative electron transport pathway and mechanism for energy conservation. The conversion of acetate to CH4 and CO2 provides only a marginal amount of energy available for ATP synthesis (DG0 = –36 kJ/CH4). A calorimetric and thermodynamic analysis of Msr. barkeri grown with acetate suggests a retarding effect of the positive enthalpy change on the driving force of growth that is overcompensated by a large positive entropy change resulting from the conversion of acetate to only gaseous products (Liu et al., 2001). Since both the enthalpy and the entropy increases are due in part to transition of CH4 and CO2 into the gaseous phase, it is proposed (Alber and Ferry, 1994) that a carbonic anhydrase (> Fig. 3, Cam) facilitates removal of CO2 from the cytoplasm by converting CO2 to membraneimpermeable HCO3 outside the cell membrane (10). þ CO2 þ H2 O ! HCO 3 þH

ð10Þ

The synthesis of Cam is up-regulated in Msr. thermophila (Alber and Ferry, 1994), Msr. acetivorans (Li et al., 2006) and Msr. mazei (Hovey et al., 2005) when switched from growth on methanol to growth on acetate consistent with a role during growth on acetate. Cam from Msr. thermophila is the archetype of an independently-evolved class of carbonic anhydrases

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(g class) and is the first carbonic anhydrase shown to function with iron in the active site (Tripp et al., 2004). The catalytic mechanism of Cam is fundamentally similar to the wellstudied a class from mammals wherein a water molecule bound to the active site metal is deprotonated yielding a metal-bound hydroxyl that attacks CO2 producing bicarbonate (Zimmerman and Ferry, 2006).

6

Research Needs

The conversion of acetate to methane by aceticlastic methanogens, is of primary importance in conversion of complex biomass to methane; thus, a comprehensive understanding of these methanogens is paramount for developing an efficient process. The enzymology of the conversion of the methyl group of acetate to methane by Methanosarcina species is fairly well understood; however, an understanding of the mechanism of energy conservation is less than adequate. Proteomic and microarray analyses have identified a plethora of proteins and genes in Methanosarcina species, many with unknown functions, that are up regulated during growth with acetate that are targets for future investigation. Genetic systems for Methanosarcina species are well established and represent a powerful tool for determining the function of cryptic genes when combined with biochemical analyses. Although Methanosaeta species are important in the conversion of acetate to methane, very little is known concerning the microbiology, biochemistry, molecular biology, and genetics. Thus, research opportunities are clearly defined that will lead to a greater understanding of aceticlastic methanogenesis.

Acknowledgments Research in the laboratory of J.G.F has been supported by the NIH, DOE, NSF, and NASA.

References Alber BE, Ferry JG (1994) A carbonic anhydrase from the archaeon Methanosarcina thermophila. Proc Nat Acad Sci USA 91: 6909–6913. Boiangiu CD et al. (2005) Sodium ion pumps and hydrogen production in glutamate fermenting anaerobic bacteria. J Mol Microbiol Biotechnol 10: 105–119. Buss KA, Cooper DR, Ingram-Smith C, Ferry JG, Sanders DA, Hasson MS (2001) Urkinase: structure of acetate kinase, a member of the ASKHA superfamily of phosphotransferases. J Bacteriol 183: 680–686. Deppenmeier U (2004) The membrane-bound electron transport system of Methanosarcina species. J Bioenerg Biomembr 36: 55–64. Eggen RIL, Geerling ACM, Boshoven ABP, Devos WM (1991) Cloning, sequence analysis, and functional expression of the acetyl coenzyme A synthetase gene from Methanothrix soehngenii in Escherichia coli. J Bacteriol 173: 6383–6389.

Ermler U, Grabarse W, Shima S, Goubeaud M, Thauer RK (1997) Crystal structure of methyl-coenzyme M reductase: the key enzyme of biological methane formation. Science 278: 1457–1462. Ferry JG (1995) CO dehydrogenase. Annu Rev Microbiol 49: 305–333. Ferry JG, Kastead KA (2007) Methanogenesis. In Archaea: Molecular Cell Biology. R Cavicchioli (ed.). Washington, DC: ASM Press, pp. 288–314. Galagan JE, et al. (2002) The genome of M. acetivorans reveals extensive metabolic and physiological diversity. Genome Res 12: 532–542. Gencic S, Grahame DA (2003) Nickel in subunit b of the acetyl-CoA decarbonylase/synthase multienzyme complex in methanogens. J Biol Chem 278: 6101–6110. Gencic S, Grahame DA (2008) Two separate one-electron steps in the reductive activation of the A cluster in

Biochemistry of Acetotrophic Methanogenesis subunit beta of the ACDS complex in Methanosarcina thermophila. Biochemistry 47: 5544–5455. Gong W, et al. (2008) Structure of the a2e2 Ni-dependent CO dehydrogenase component of the Methanosarcina barkeri acetyl-CoA decarbonylase/synthase complex. Proc Natl Acad Sci USA 105: 9558–9563. Gorrell A, Ferry JG (2007) Investigation of the Methanosarcina thermophila acetate kinase mechanism by fluorescence quenching. Biochemistry 46: 14170–14176. Gottschalk G, Thauer RK (2001) The Na+translocating methyltransferase complex from methanogenic archaea. Biochim Biophys Acta 1505: 28–36. Gu WW, Gencic S, Cramer SP, Grahame DA (2003) The A-cluster in subunit beta of the acetyl-CoA decarbonylase/synthase complex from Methanosarcina thermophila: Ni and Fe K-Edge XANES and EXAFS analyses. J Am Chem Soc 125: 15343–15351. Hedderich R, Hamann N, Bennati M (2005) Heterodisulfide reductase from methanogenic archaea: a new catalytic role for an iron-sulfur cluster. Biol Chem 386: 961–970. Hovey R, et al. (2005) DNA microarray analysis of Methanosarcina mazei Go1 reveals adaptation to different methanogenic substrates. Mol Genet Genomics 273: 225–239. Latimer MT, Painter MH, Ferry JG (1996) Characterization of an iron-sulfur flavoprotein from Methanosarcina thermophila. J Biol Chem 271: 24023–24028. Lawrence SH, Luther KB, Schindelin H, Ferry JG (2006) Structural and functional studies suggest a catalytic mechanism for the phosphotransacetylase from Methanosarcina thermophila. J Bacteriol 188: 1143–1154. Li L, et al. (2007) Quantitative proteomic and microarray analysis of the archaeon Methanosarcina acetivorans grown with acetate versus methanol. J Proteome Res 6: 759–771. Li Q, Li L, Rejtar T, Lessner DJ, Karger BL, Ferry JG (2006) Electron transport in the pathway of acetate conversion to methane in the marine archaeon Methanosarcina acetivorans. J Bacteriol 188: 702–710. Liu JS, Marison IW, von Stockar U (2001) Microbial growth by a net heat up-take: A calorimetric and thermodynamic study on acetotrophic methanogenesis

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by Methanosarcina barkeri. Biotechnol Bioeng 75: 170–180. Maupin-Furlow J, Ferry JG (1996) Characterization of the cdhD and cdhE genes encoding subunits of the corrinoid iron-sulfur enzyme of the CO dehydrogenase complex from Methanosarcina thermophila. J Bacteriol 178: 340–346. Meuer J, Kuettner HC, Zhang JK, Hedderich R, Metcalf WW (2002) Genetic analysis of the archaeon Methanosarcina barkeri Fusaro reveals a central role for Ech hydrogenase and ferredoxin in methanogenesis and carbon fixation. Proc Natl Acad Sci USA 99: 5632–5637. Murakami E, Deppenmeier U, Ragsdale SW (2001) Characterization of the intramolecular electron transfer pathway from 2-hydroxyphenazine to the heterodisulfide reductase from Methanosarcina thermophila. J Biol Chem 276: 2432–2439. Nelson MJK, Ferry JG (1984) Carbon monoxidedependent methyl coenzyme M methylreductase in acetotrophic Methanosarcina spp. J Bacteriol 160: 526–532. Pisa KY, Weidner C, Maischak H, Kavermann H, Muller V (2007) The coupling ion in the methanoarchaeal ATP synthases: H+versus Na+in the A1Ao ATP synthase from the archaeon Methanosarcina mazei Go1. FEMS Microbiol Lett 277: 56–63. Ragsdale SW (2004) Life with carbon monoxide. Crit Rev Biochem Mol Biol 39: 165–195. Simianu M, Murakami E, Brewer JM, Ragsdale SW (1998) Purification and properties of the hemeand iron-sulfur-containing heterodisulfide reductase from Methanosarcina thermophila. Biochemistry 37: 10027–10039. Smith KS, Ingram-Smith C (2007) Methanosaeta, the forgotten methanogen? Trends Microbiol 7: 150–155. Spector LB (1980) Acetate kinase: a triple displacement enzyme. Proc Natl Acad Sci USA 77: 2626–2630. Thauer RK, Seigo S (2007) Methane as fuel for anaerobic microorganisms. Ann NY Acad Sci 1125: 158–170. Tripp BC, Bell CB, Cruz F, Krebs C, Ferry JG (2004) A role for iron in an ancient carbonic anhydrase. J Biol Chem 279: 6683–6687. Zimmerman SA, Ferry JG (2006) Proposal for a hydrogen bond network in the active site of the prototypic g-class carbonic anhydrase. Biochemistry 45: 5149–5157.

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24 Aliphatic Hydrocarbons, Carbon–Carbon Bond Formation L. P. Wackett Department of Biochemistry, Molecular Biology, and Biophysics and BioTechnology Institute, University of Minnesota, St. Paul, MN, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 370 2 Types of Carbon–Carbon Bond Forming Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 370 3 Alkanes from Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 371 4 Alkene Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372 5 Cyclic Aliphatic Rings . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 372 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 373

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Abstract: Biologically produced, non-gaseous aliphatic hydrocarbons typically require carbon–carbon bond formation steps to elaborate the hydrocarbon chains. A major source of biological aliphatic hydrocarbon chains are fatty acids. Every other carbon–carbon bond in a fatty acid chain derives from a Claisen condensation reaction. In fatty acid biosynthesis, a decarboxylative Claisen condensation mechanism is most prevalent. The chains may be decarbonylated to produce alkanes, or condensed to produce alkenes. Fatty acid modification reactions can also occur via novel carbon–carbon bond forming mechanisms.

1

Introduction

Microbes biosynthesize hydrocarbons: alkanes, alkenes, arenes, and isoprenoid compounds. In many cases, the biosynthetic pathways have not been fully elucidated. Currently, there is a growing interest to use these metabolic pathways to generate molecules that can be used as fuels or specialty chemicals. The goal would be to engineer microorganisms that transform biomass sugars to hydrocarbons that could be used in place of petroleum-based fuels or chemicals. In general, to go from sugar based molecules with a general chemical formula (CHO)x to a hydrocarbon, of the general chemical formula (CH2)x, one needs to remove oxygen atoms and input electrons and protons. If the hydrocarbon is greater in length than a typical C5 or C6 sugar molecule, then carbon–carbon bond forming reactions are needed. Moreover, the hydrocarbons may not be made directly from sugars but rather may proceed via an acetate (C2) intermediate. This requires the generation of numerous carbon–carbon bonds to build up the hydrocarbon chains.

2

Types of Carbon–Carbon Bond Forming Reactions

Important carbon–carbon bond forming reactions are the Claisen condensation and an aldolase-type mechanism (Heath and Rock, 2002). The former is perhaps more prevalent in biology for the formation of hydrocarbons and hydrocarbon fragments and will be the focus of the discussion here. The Claisen condensation in biology is best represented by the thiolasecatalyzed reaction shown in > Fig. 1. In this reaction, a carbon–carbon bond is formed with the concomitant cleavage of the thioester carbonyl carbon-to-sulfur bond. The Claisen condensation figures prominently in the synthesis of fatty acids, polyketides, terpenes, and steroids. There is a second type of Claisen condensation reactions in which one reactant is an acylCoA and the other is the a-carboxylate of an alkanoic acid (> Fig. 2). In this reaction,

. Figure 1 Thiolase catalyzed reaction can occur to the right (biosynthetic) or the left (degradative) to make or break a carbon–carbon bond, respectively.

Aliphatic Hydrocarbons, Carbon–Carbon Bond Formation

24

. Figure 2 Decarboxylative Claisen condensation in which one carbon-to-carbon bond is broken and another is formed.

decarboxylation generates a transient carbanion that can attack the carbonyl carbon of an acylCoA. The result is the formation of a new carbon-to-carbon bond with the displacement of the CoA thiolate. This is the major carbon chain-building reaction in the biosynthesis of fatty acids in microbes, plants, and animals. The decarboxylative Claisen condensation in fatty acid biosynthesis (> Fig. 2) uses an activated form of the acetyl unit that is ultimately condensed to elongate a fatty acid chain two carbons at a time. The activation step requires an investment of ATP to form malonyl-CoA. This serves to provide the energy that ultimately drives carbon–carbon bond formation. While the non-decarboxylative Claisen condensation (> Fig. 1) can form carbon-to-carbon bonds, the equilibrium is largely in the biodegradative direction. This is probably why cells have evolved to use the decarboxylative mechanism. It drives the reactions in favor of biosynthesis when the cells need fatty acids for membranes during rapid cell division.

3

Alkanes from Fatty Acids

Fatty acids, or their close relatives, are the obvious source of carbon for long chain, nonisoprenoid hydrocarbons in biological systems. Alkyl hydrocarbons are produced by plants and animals for purposes of water repulsion or as solvents. For example, some plants produce hydrocarbons to coat leaf surfaces to prevent dessication. Insects make various toxic sprays for defense, some of which contain alkanes. The reaction(s) generating alkanes are not elucidated in detail. However, there is evidence that fatty aldehydes can undergo a decarbonylation reaction that generates the corresponding n-1 alkane and carbon monoxide. In one study, the decarbonylase is proposed to be a cobalt porphyrin enzyme (Dennis and Kolattukudy, 1992). In other cases, a cytochrome P450 heme-iron monooxygenase is reported to catalyze decarbonylation reactions (Coon and Vaz, 1988). A novel mechanism for alkane formation was recently proposed to occur in the bacterium Vibrio furnissii M1 (Park et al., 2005). This strain was reported to reduce fatty acids to the corresponding alcohol with subsequent reduction of the alcohol to the alkane (Park, 2005). Thus, there was no net loss of carbon in the reaction sequence generating the alkane. Alcohol to alkyl carbon reduction reactions are rare in biology. It is best exemplified by the reduction of a ribonucleotide to a deoxyribonucleotide by ribonucleotide reductase (Nordlund and Reichard, 2006). Ribonucleotide reductases from different organisms contain different metal cofactors. The metal cofactors serve to generate a carbon-centered radical intermediate during the reduction of the carbon–oxygen bond of the hydroxyl group on the ribose sugar. The radical is then further reduced leading to an overall two electron reduction of an alcohol to a methylene carbon. There is evidence for radical stabilization by resonance between C2 and C3

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on the sugar moiety. The reaction proposed to be catalyzed by V. furnissii M1 would have had to be even more powerful, to reduce a completely unactivated alkyl alcohol and thus generate a putatively unstabilized carbon-centered radical. In an effort to study this novel biological alcohol reduction reaction, V. furnissii M1 was studied via studies on whole cell physiology, cell-free enzyme activity and genomics (Wackett et al., 2007). The latter approach was designed to identify genes and thus allow subsequent metabolic engineering for developing recombinant alkane-synthesizing strains. However, no putative alkane-biosynthetic genes could be identified. Moreover, no genes for alkane oxidation could be discerned. This was surprising as bacteria that make and store highly reduced forms of carbon typically have genes that encode the catabolism of those compounds. In cell physiological studies, no alkanes could be discerned after solvents and workup procedures were optimized to eliminate contaminating alkanes. Cell-free studies were then conducted and showed no evidence for alcohol reductase activity. It was concluded that alkane formation by V. furnissii M1 was not reproducible.

4

Alkene Biosynthesis

Alkenes are also found in biological systems; for example, plants make 1-alkenes (Gorgen and Boland, 1989). The enzymes underlying 1-alkene biosynthesis have not been identified but labeling studies have provided insights into the mechanism. There is evidence for an oxidative decarboxylation of fatty acids to generate 1-alkenes. Phenylalkyl acids were also shown to undergo this reaction in feeding studies. Another different class of alkenes are internal alkenes that are made by some gram positive bacterial genera (Tornabene, 1980). The natural products made are long chain, generally C23–C31 alkenes, with the double bond near the center of the molecule. Radiolabeling studies are consistent with the alkenes being derived from two fatty acid precursor molecules. The reaction(s) has been described as a head-to-head fatty acid condensation. The mechanism of the condensation has yet to be elucidated. It has been established that the carboxylate carbon atoms from one of the fatty acids is displaced during the condensation. The double bond is proposed to have a cis-relative stereochemistry but this remains to be firmly established. Overall, the biological function for median alkene biosynthesis also remains to be elucidated. Alkenes appear to be made constitutively, so there occurrence has not been linked to any specific growth condition or cellular need.

5

Cyclic Aliphatic Rings

The enzyme cyclopropane fatty acid synthase catalyzes methyl group transfer from S-adenosyl-L-methionine to the double bond of unsaturated fatty acids to make a cyclopropane ring (Iwig et al., 2004). Fatty acids containing a cyclopropane ring are found in divergent bacterial genera, typically being present more in late exponential and stationary growth phases. An even more unusual cycloalkyl biological structure is the linearly fused cyclobutane rings denoted as ladderanes (Sinninghe Damste´ et al., 2002). The ladderane lipids have been found in slow-growing anaerobic bacteria that catalyze anaerobic ammonia oxidation, the so-called anammox reactions.

Aliphatic Hydrocarbons, Carbon–Carbon Bond Formation

6

24

Research Needs

In the petroleum era, preformed carbon–carbon bonds were plentiful. Oil extraction provided aromatic ring carbon structures, and long carbon–carbon aliphatic chains. The energy inherent in these bonds could thus be used in reformulation to produce an enormous array of organic structures for society. A new industrial era based on biofuels will usher in a new age of interest in carbon–carbon bond formation. Biological systems most commonly break down carbon chains into small, common precursors and build up most molecules de novo. Several major classes of carbon–carbon bond making biochemistry have been elucidated over the last century. However, there is clearly a need for new discovery. Additionally, currently known enzymes can be used in new ways with surrogate substrates, or following enzyme engineering, to build new molecules that are not typically formed in biological systems. This will be an increasingly important field of research.

References Coon MJ, Vaz AD (1988) Role of cytochrome P-450 in hydrocarbon formation from xenobiotic and lipid hydroperoxides. Prog Clin Biol Res 274: 497–507. Dennis M, Kolattukudy PE (1992) A cobalt-porphyrin enzyme converts a fatty aldehyde to a hydrocarbon and CO. Proc Natl Acad Sci, USA 89: 5306–5310. Gorgen G, Boland W (1989) Biosynthesis of 1-alkenes in higher plants: stereochemical implications. A model study with Carthamus tinctorius (Asteraceae). Eur J Biochem 185: 237–242. Heath RJ, Rock CO (2002) The Claisen condensation in biology. Nat Prod Rev 19: 581–596. Iwig DF, Grippe AT, McIntyre TA, Booker SJ (2004) Isotope and elemental effects indicate a rate-limiting methyl transfer as the initial step in the reaction catalyzed by Escherichia coli cyclopropane fatty acid synthase. Biochemistry 43: 13510–13524. Nordlund P, Reichard P (2006) Ribonucleotide reductases. Annu Rev Biochem 75: 681–706.

Park MO (2005) New pathway for long-chain n-alkane synthesis via 1-alcohol in Vibrio furnissii M1. J Bacteriol 187: 1426–1429. Park MO, Heguri K, Hirata K, Miyamoto K (2005) Production of alternatives to fuel oil from organic waste by the alkane-producing bacterium, Vibrio furnissii M1. J Appl Microbiol 98: 324–331. Sinninghe Damste´ JS, Strous M, Rijpstra WI, Hopmans EC, Geenevasen JA, van Duin AC, van Niftrik LA, Jetten MS (2002) Linearly concatenated cyclobutane lipids form a dense bacterial membrane. Nature 419: 708–712. Tornabene TG (1980) Formation of hydrocarbons by bacteria and algae. Basic Life Sci 18: 421–438. Wackett LP, Frias J, Seffernick JL, Sukovich D, Cameron SM (2007) Vibrio furnissii M1: Genomic and biochemical studies demonstrating the absence of an alkane-producing phenotype. Appl Environ Microbiol 73: 7192–7198.

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25 Halogenated Organic Compounds – CarbonHalogen Bond Formation C. D. Murphy School of Biomolecular and Biomedical Science and Centre for Synthesis and Chemical Biology, University College Dublin, Belfield, Dublin, Ireland [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376

2 2.1 2.2 2.3 2.4

Biohalogenation Mechanisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 Haloperoxidases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 376 FADH2-Dependent Halogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 377 Non-Heme Fe (II), O2- and a-Ketoglutarate Dependent Halogenases . . . . . . . . . . . . . . 379 SAM-Dependent Halogenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 380

3

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 381

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Halogenated Organic Compounds – Carbon-Halogen Bond Formation

Abstract: Microorganisms are significant producers of halogenated natural products. Our understanding of the enzymatic mechanisms responsible for the incorporation of halogens into biochemical intermediates has changed dramatically in the last decade, and has virtually excluded involvement of haloperoxidases. Electron-rich substrates such as phenyl and pyrrole are typically halogenated via FADH2-dependent halogenases that produce hyphohalous acid as the initial halogenating species. Unactivated carbon centers are halogenated by non-heme Fe(II), a-ketoglutarate and O2-dependent enzymes, which are related to Fe (II)/a-ketoglutarate hydroxylases. Enzymes that catalyze nucleophilic chlorination are much less common; those that have been observed employ the substrate S-adenosyl methionine, which is also the substrate for the only wild-type fluorinating enzyme described to date.

1

Introduction

Microorganisms produce a highly diverse collection of halogenated natural products as pigments, antibiotics, signaling molecules and biochemical intermediates. These compounds are of considerable interest owing to their biological activity, and the biochemical mechanisms involved in their production are intriguing. Microorganisms have been known to be a source of halogenated compounds for over 100 years, although it was thought for quite some time that these compounds occurred infrequently in nature; by 2004, over 4000 halometabolites had been identified (Gribble, 2004). Fluorinated and iodinated natural products are relatively rare; only three microbial fluorinated natural products are known from two species of Streptomyces. Algae are known to produce iodometabolites, such as iodomethane (Murphy et al., 2000), and iodinated nucleosides (Kazlauskas et al., 1983). Terrestrial bacteria and fungi are important sources of chlorinated metabolites, while a diverse array of organobromine compounds including alkanes, terpenes, polyketides and peptides have been isolated from marine microorganisms (Gribble, 1999). The enzymology of halogenation reactions in microorganisms is a highly active research topic for several reasons. Aside from the fundamental interest in how halogen atoms are specifically incorporated into biological compounds, the enzymes responsible have biotechnological potential as alternatives to classical organic methods for halogenation reactions. Also, the presence of halogens in a secondary metabolite can dramatically alter its biological properties, thus knowledge of the biosynthetic mechanism and genes involved in the production of halometabolites can lead to the development of novel antibiotics.

2

Biohalogenation Mechanisms

2.1

Haloperoxidases

The most well studied class of halogenating enzyme is the haloperoxidase, which catalyze the hydrogen-peroxide dependent oxidation of chloride, bromide or iodide ions (Neidleman and Geigert, 1986). The enzymes are classified depending on (i) the range of halides that are oxidized, and (ii) the nature of the prosthetic group. Chloroperoxidases oxidize Cl
, Br
and I
, bromoperoxidases oxidize Br
and I
, and iodoperoxidases oxidize I
only. Haloperoxidases with ferriprotoporphyrin IX or vanadium as the prosthetic group have been identified. Perhydrolases that do not contain either heme or vanadium, but catalyze halogenation

Halogenated Organic Compounds – Carbon-Halogen Bond Formation

25

. Figure 1 Haloperoxidase-catalysed chlorination of monochlorodimedone.

reactions, are not classified as haloperoxidases since the oxidation of halide is independent of the enzyme, which catalyses the H2O2-dependent oxidation of short chain carboxylic acids to peracids, which in turn spontaneously oxidize halide ion (van Pee, 2001). The chloroperoxidase from Caldariomyces fumago was the first haloperoxidase to be isolated during investigations on the biosynthesis of caldariomycin (Morris and Hager, 1966). The spectrophotometric assay developed in these studies, which employed the synthetic substrate monochlorodimedone (> Fig. 1), was subsequently applied to the identification of similar enzymes in a range of organisms. In almost all of these studies no regard was paid to the natural substrate for the proposed halogenation reaction, and it was assumed that haloperoxidases were responsible for the biosynthesis of most halometabolites. However, haloperoxidases display very broad substrate specificity and no regioselectivity, which is not consistent with the likely biosynthetic mechanisms of most naturally produced organohalogens (vanPee, 1996). Furthermore, disruption of genes encoding haloperoxidases did not result in the elimination of production of halometabolites (Kirner et al., 1996), and when the gene cluster for chlortetracycline biosynthesis in S. aureofaciens was sequenced, none was homologous to those known to encode haloperoxidases (Dairi et al., 1995). X-ray crystallography of the C. fumago enzyme revealed that hypohalous acid is produced via oxidation of the ferriprotoporphyrin IX, yielding an oxyferryl centre that in turn oxidizes halide ion (Sundaramoorthy et al., 1998). At acidic pH, which is the optimum for most haloperoxidasecatalyzed halogenation reactions, hypohalous acid is generated. No substrate-binding site could be detected, which is consistent with the inability to determine Michaelis constant values for the organic substrate, thus the hypohalous acid is released from the active site where is reacts spontaneously with electron-rich organic compounds. The function of haloperoxidases is still under investigation, and it has been speculated that in marine algae, such as Laminaria digitata, iodoperoxidase might be involved in the uptake of iodine (Colin et al., 2005).

2.2

FADH2-Dependent Halogenases

The realization that haloperoxidases were not involved in the biosynthesis of most halometabolites prompted the search for other halogenating enzymes. In 1997 a new halogenating enzyme was discovered in cell extracts of Pseudomonas fluorescens expressing the gene prnA, which coded for the enzyme responsible for chlorination of tryptophan in pyrrolnitrin biosynthesis (Hohaus et al., 1997). Purification of the enzyme revealed that it required

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O2 for activity and FADH2 as a co-factor, and the presence of an NADH-dependent reductase to reduce FAD (Keller et al., 2000). As gene clusters responsible for the biosynthesis of halometabolites have been sequenced, other FADH2-dependent halogenases have been identified based on sequence homology. A relatively small number of FADH2-dependent halogenases have been studied in vitro, because in many cases the substrate for the enzyme is not known or is difficult to synthesize. For example, PltA, which is the halogenase involved in pyoluteroin biosynthesis in P. fluorescens Pf5, catalyses the dichlorination of a pyrrole moiety bound to a peptidyl carrier protein, PltL (Dorrestein et al., 2005). X-ray crystallographic analysis of PrnA shed light on the mechanism of regiospecific chlorination (Dong et al., 2005). A flavin hydroperoxide is formed by oxidation of FADH2 by oxygen, and subsequent attack on the isoalloxazine by Cl
yields HOCl, which travels along a 10 A˚ tunnel to the tryptophan binding site. Lysine 79, which is located in the tunnel, was identified as having a possible role in the mechanism of chlorination, and a K79A mutant had no chlorinating activity. A glutamate residue in the tryptophan binding site was also identified as playing a key role in the reaction, since an E346Q mutant resulted in a significantly reduced kcat compared with the wild type enzyme. Dong et al proposed that K79 activates the HOCl via hydrogen bonding (> Fig. 2a) and E346 stabilizes a putative Wheland intermediate. The crystal structure of another tryptophan 7-halogenase, RebH, which is involved in rebeccamycin biosynthesis, has been solved and has 54% identity with PrnA (Yeh et al., 2007). Furthermore, the K79 is conserved in RebH and mutation of this residue results in an inactive enzyme. Experiments with radioactive chlorine-36 indicated that chlorine was bound to the enzyme in the absence of tryptophan, and when the enzyme was subsequently eluted from a gel filtration column it was still possible to generate chlorotryptophan upon addition of tryprophan. Thus, it was suggested that a stable enzyme-bound chlorinating species was involved in the mechanism of chlorination, and

. Figure 2 FADH2-dependent chlorination of tryptophan via HOCl (a) and lysine chloramine (b).

Halogenated Organic Compounds – Carbon-Halogen Bond Formation

25

this could be a chloramine formed by reaction of HOCl and the e-NH2 of K79 (> Fig. 2b). However, as yet there is no direct evidence of this species. Gene sequences of FADH2-dependent halogenases reveal that there are regions of conservation: the FAD binding site has a GxGxxG motif common to nucleotide binding regions in other enzymes, and a WxWxIP motif, which is absolutely conserved in all halogenases of this type (van Pee and Patallo, 2006). The trp residues are located close to the flavin and probably prevent substrate interaction with flavin hydroperoxide as would occur in monooxygenases.

2.3

Non-Heme Fe (II), O2- and a-Ketoglutarate Dependent Halogenases

Unactivated carbon centres cannot be halogenated via FADH2-dependent halogenases, and the biosynthesis of compounds containing one or more halogen atoms on a methyl or methylene involve a different class of halogenating enzyme: the non-heme Fe (II), a-ketoglutarate and O2-dependent halogenase. The best studied is of this class SyrB2, which chlorinates L-threonyl-SyrB1 (peptidyl carrier protein) to yield the 4-chlorothreonine residue in syringomycin (Vaillancourt et al., 2005), which is a lipodepsipeptide produced by P. syringae. Crystallographic analysis revealed that this enzyme is structurally similar to Fe (II)/a-ketoglutarate hydroxylases, which typically have a facial triad of two histidine residues and a glutamate or aspartate that co-ordinate the iron (Blasiak et al., 2006). However, while SyrB2 has the histidine residues, there is no glutamate/ aspartate, instead there is an alanine (Ala 118, conserved in mononuclear iron halogenases), which creates space for chloride ion to occupy and co-ordinate with the iron. When this happens a conformational change occurs that closes the active site and limits access to substrates, via a 17 A˚ tunnel. The peptidyl carrier protein SyrB1 presents threonine attached to the phosphopantetheine cofactor, to the active site of SyrB2. The interaction of the substrate and enzyme also excludes water from the active site and allows O2 to bind. The most likely mechanism of halogenation is shown in > Fig. 3, and proceeds with the generation of a substrate carbon radical after decarboxylation of a-ketoglutarate and formation of a ferryl-oxo intermediate, which abstracts a hydrogen atom from the substrate. Abstraction of the chlorine atom by the substrate radical would yield the 4-chloro-L-threonyl-SyrB1 and return the iron to the + 2 state. Spectroscopic evidence for the presence of Fe(IV) intermediates in this class of halogenase was reported by Galonic et al. (2007). By employing three proteins involved in cytotrienin biosynthesis in Streptomyces sp, CytC1 (adenylation), CytC2 (thiolation) and CytC3 (halogenation), which together catalyze the halogenation of L-aminobutyric acid (LAba), these researchers demonstrated the presence of intermediates in the catalytic cycle via absorption spectroscopy. The initial (anaerobic) CytC3-a-KG-Cl
-L-Aba-S-CytC2 complex absorbs maximally at 520 nm. Addition of oxygen starts the reaction and an intermediate can be detected that absorbs maximally at 318 nm characteristic of a Fe (IV)-oxo intermediate, and which subsequently decays in concert with the starting complex. The lifetime of the intermediate could be prolonged using deuterated aminobutyric acid as the substrate, and this large kinetic isotope effect indicated that it is involved in abstraction of hydrogen from the substrate. Mo¨ssbauer spectroscopy detected two species that are characteristic of Fe(IV)-oxo intermediates that have been observed in a-KG-dependent oxygenases, such as taurine dioxygenase, further supporting the hypothesis that the mechanisms of mononuclear Fe(II) hydroxylases and halogenases are analogous.

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. Figure 3 Mechanism of non-heme Fe (II), O2 and a-ketoglutarate-dependent halogenases (R = –CH2–C–CH(NH2)–CO–S–CO2; R1 = –(CH2)2–CO2–). Reprinted by permission from Macmillan Publishers Ltd: Nature Chemical Biology 3, 113-116, copyright 2008.

2.4

SAM-Dependent Halogenases

Methyl transferases that catalyse the production of halomethanes from S-adenosyl methionine (SAM) and halide ion have been isolated from fungi, bacteria and plants. The biogenesis of these compounds is not clear, but it has been suggested that in salt-tolerant plants, such as Batis maritima, emission of chloromethane might be a mechanism of regulating halide ion concentrations (Ni and Hager, 1999). Chloromethane is a methyl donor in veratryl alcohol biosynthesis in wood rotting fungi, and its emission is possibly linked to the uncoupling of metabolism in senescence (Harper, 2000). Recently two new halogenating enzymes that employ SAM as a substrate have been discovered. The bacterium Streptomyces cattleya produces the rare fluorinated natural products fluoroacetate and 4-fluorothreonine. The fluorinase (FlA) responsible for the formation of the C-F bond catalyses the nucleophilic displacement of methionine from SAM by fluoride (O’Hagan et al., 2002), yielding 50 -fluoro-50 -deoxyadenosine, which is subsequently transformed to the

Halogenated Organic Compounds – Carbon-Halogen Bond Formation

25

. Figure 4 SAM-dependent halogenation (X = Cl/F).

common biosynthetic precursor of fluoroacetate and 4-fluorothreonine, fluoroacetaldehyde (> Fig. 4). Crystallographic analysis and site directed mutagenesis of FlA (Zhu et al., 2007) revealed that heavily hydrated fluoride ion binds weakly in the active site and the water molecules are replaced by hydrogen bonds to the amide NH and side chain OH of an active site serine (S158). The subsequent binding of SAM results in final desolvation of the fluoride ion, making it a strong nucelophile that attacks the C-5’ of SAM. From analysis of the sal biosynthetic gene cluster, responsible for the production of the non-ribosomal peptide/polyketide hybrid salinosporamide in the marine bacterium Salinispora tropica, salL emerged as the gene likely to be involved in halogenation (Eustaquio et al., 2008). SalL has a 35% amino acid identity with FlA, and subsequent in vitro experiments demonstrated that the enzyme catalyses the production of 5’chloro-5’deoxyadenosine from SAM and Cl
(> Fig. 4). The enzyme can also incorporate bromine and iodine, but, significantly, not fluorine. X-ray crystallographic analysis of SalL revealed that the two enzymes share a similar architecture, but there are some critical differences, most notably the S158 in FlA is replaced by a glycine in SalL, which provides a larger pocket for halide binding, and a 23-residue loop is missing from the N-terminal domain of SalL, which in FlA compresses the loop carrying S158, bringing it into contact with the ribose unit of the substrate.

3

Research Needs

The discoveries of FADH2-dependent halogenases, non-heme Fe(II)/a-ketoglutaratedependent halogenases and SAM dependent chlorinase and fluorinase has transformed our understanding of biohalogenation. Undoubtedly more halogenases remain to be discovered,

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and sequencing of biosynthetic gene clusters will play an invaluable role in this; the more difficult aspect seems to be the successful observation of the halogenation reaction in vitro. The mechanisms of the current known enzymes have been made clear by x-ray crystallography, although, in the case of FADH2-dependent halogenases, the precise role of HOCl in the halogenating reaction has yet to emerge. A greater knowledge of these fascinating enzymes may lead to their application as biocatalysts, or in the generation of novel halogenated compounds, such as antibiotics, via engineering of biosynthetic gene clusters (See > Chapter 37, Vol. 4, Part 5).

References Blasiak LC, Vaillancourt FH, Walsh CT, Drennan CL (2006) Crystal structure of the non-haem iron halogenase SyrB2 in syringomycin biosynthesis. Nature 440: 368–371. Colin C et al. (2005) Vanadium-dependent iodoperoxidases in Laminaria digitata, a novel biochemical function diverging from brown algal bromoperoxidases. J Biol Inorg Chem 10: 156–166. Dairi T, Nakano T, Aisaka K, Katsumata R, Hasegawa M (1995) Cloning and nucleotide sequence of the gene responsible for chlorination of tetracycline. Biosci Biotechnol Biochem 59: 1099–1106. Dong CJ, Flecks S, Unversucht S, Haupt C, van Pee KH, Naismith JH (2005) Tryptophan 7-halogenase (PrnA) structure suggests a mechanism for regioselective chlorination. Science 309: 2216–2219. Dorrestein PC, Yeh E, Garneau-Tsodikova S, Kelleher NL, Walsh CT (2005) Dichlorination of a pyrrolyl-Scarrier protein by FADH2-dependent halogenase PltA during pyoluteorin biosynthesis. In Proceedings of the National Academy of Sciences of the United States of America, Vol. 102, pp. 13843–13848. Eustaquio AS, Pojer F, Noe JP, Moore BS (2008) Discovery and characterization of a marine bacterial SAMdependent chlorinase. Nat Chem Biol 4: 69–74. Galonic DP, Barr EW, Walsh CT, Bollinger JM, Krebs C (2007) Two interconverting Fe((IV)) intermediates in aliphatic chlorination by the halogenase CytC3. Nat Chem Biol 3: 113–116. Gribble GW (1999) The diversity of naturally occurring organobromine compounds. Chem Soc Rev 28: 335–346. Gribble GW (2004) Natural organohalogens: a new frontier for medicinal agents? J Chem Educ 81: 1441–1449. Harper DB (2000) The global chloromethane cycle: biosynthesis, biodegradation and metabolic role. Nat Prod Rep 17: 337–348. Hohaus K et al. (1997) NADH-dependent halogenases are more likely to be involved in halometabolite

biosynthesis than haloperoxidases. Angew Chem Int Ed (English) 36: 2012–2013. Kazlauskas R, Murphy PT, Wells RJ, Bairdlambert JA, Jamieson DD (1983) Halogenated pyrrolo[2,3-D] pyrimidine nucleosides from marine organisms. Aust J Chem 36: 165–170. Keller S, Wage T, Hohaus K, Holzer M, Eichhorn E, van Pee KH (2000) Purification and partial characterization of tryptophan 7-halogenase (PrnA) from Pseudomonas fluorescens. Angew Chem Int Ed (English) 39: 2300–2302. Kirner S, Krauss S, Sury G, Lam ST, Ligon JM, vanPee KH (1996) The non-haem chloroperoxidase from Pseudomonas fluorescens and its relationship to pyrrolnitrin biosynthesis. Microbiology 142: 2129–2135. Morris DR, Hager LP (1966) Chloroperoxidase.I. Isolation and properties of crystalline glycoprotein. J Biol Chem 241: 1763–1738. Murphy CD, Moore RM, White RL (2000) An isotopic labeling method for determining production of volatile organohalogens by marine microalgae. Limnol Oceanogr 45: 1868–1871. Neidleman SL, Geigert J (1986) Biohalogenation: Principles, Basic Roles and Applications. Chichester: Ellis Horwood. Ni XH, Hager LP (1999) Expression of Batis maritima methyl chloride transferase in Escherichia coli. Proc Nat Acad Sci USA 96: 3611–3615. O’Hagan D, Schaffrath C, Cobb SL, Hamilton JTG, Murphy CD (2002) Biosynthesis of an organofluorine molecule – A fluorinase enzyme has been discovered that catalyses carbon-fluorine bond formation. Nature 416: 279–279. Sundaramoorthy M, Terner J, Poulos TL (1998) Stereochemistry of the chloroperoxidase active site: crystallographic and molecular-modeling studies. Chem Biol 5: 461–473. Vaillancourt FH, Yin J, Walsh CT (2005) SyrB2 in syringomycin E biosynthesis is a nonheme Fe-II

Halogenated Organic Compounds – Carbon-Halogen Bond Formation a-ketoglutarate- and O2-dependent halogenase. Proc Nat Acad Sci USA 102: 10111–10116. van Pee KH (2001) Microbial biosynthesis of halometabolites. Arch Microbiol 175: 250–258. van Pee KH, Patallo EP (2006) Flavin-dependent halogenases involved in secondary metabolism in bacteria. Appl Microbiol Biotechnol 70: 631–641. Van Pee KH (1996) Biosynthesis of halogenated metabolites by bacteria. Annu Rev Microbiol 50: 375–399.

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Yeh E, Blasiak LC, Koglin A, Drennan CL, Walsh CT (2007) Chlorination by a long-lived intermediate in the mechanism of flavin-dependent halogenases. Biochemistry 46: 1284–1292. Zhu XF, Robinson DA, McEwan AR, O’Hagan D, Naismith JH (2007) Mechanism of enzymatic fluorination in Streptomyces cattleya. J Am Chem Soc 129: 14597–14604.

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26 Formation of Fatty Acids I. M. Lo´pez-Lara* . O. Geiger Centro de Ciencias Geno´micas, Universidad Nacional Auto´noma de Me´xico, Cuernavaca, Morelos, Mexico *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 386

2 Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 387 2.1 The Basic Type II Fatty Acid Biosynthesis Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 388 2.2 Modifications of Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 389 3

Fatty Acids and Fatty Acid-Derivatives as Signal Molecules . . . . . . . . . . . . . . . . . . . . . 390

4

Specificity of Acyltransferases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 391

5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 392

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_26, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Fatty acids are the building blocks of diverse membrane lipids and therefore are essential for the viability of all cells, except the Archaea where side chains of membrane lipids are isoprenoids. There are multiple examples of fatty acid-derived molecules as signal compounds for communication of cell-to-cell among prokaryotes in one species, interspecies, or even interkingdom (i.e., between prokaryotes and eukaryotes). Fatty acid biosynthesis is catalyzed in most bacteria by a group of highly conserved proteins known as the type II fatty acid synthase (FASII) system. A key protein in this system is the acyl carrier protein (ACP) which acts as carrier of growing acyl chains during biosynthesis and as donor of acyl chains during transfer to target molecules. The Escherichia coli FAS II system is the model pathway of fatty acid biosynthesis in bacteria and mainly the same set of enzymes is present in different bacteria. Variations of the basic set give rise to the diversity of fatty acid composition of each organism. The intrinsic specificity of the acyltransferases is largely responsible for the fatty acid composition of complex lipids and signals molecules.

1

Introduction

Fatty acids (FAs) are organic molecules consisting of an aliphatic carbon chain with a methyl group at one end of the chain and a carboxylic acid group (carbon number 1) at the other end. Fatty acids are key components of the cell, and their synthesis is essential for all organisms except for Archaea, although the presence of fatty acids as part of lipoproteins has been described in Archaea (Pugh and Kates, 1994). FAs occur in their free form only in small amounts; in most cells they are built-up to form complex lipids with structural, energy storage, and signalling functions. In glycerophospholipids, which are usually the major components of cell membranes, FAs are ester-linked to a glycerol backbone, although there are many other membrane-forming lipids which are based on FAs (Geiger et al., 2009). FAs are also constituents of the lipid A part of lipopolysaccharides which are constituents of the outer membrane of Gram-negative bacteria (review by Raetz et al., 2007, Lipid A chapter 28), and of lipoteichoic acids that contribute to the cell wall of Gram-positive bacteria (Neuhaus and Baddiley, 2003). They are also used for posttranslational protein modifications that are functionally important (e.g., Rezwan et al., 2007). Triacylglycerols are FA triesters of glycerol widely used as reserve compounds among Eukarya including yeast and fungi, however, the occurrence of triacylglycerols in bacteria seems to be restricted to the Actinomycetes group (Alvarez and Steinbu¨chel, 2002). Instead, most bacteria synthesize polymeric lipids such as poly(3-hydroxybutyrate) (PHB) or other polyhydroxyalkanoates (PHAs) as reserve material (Steinbu¨chel, 2001). In many cases fatty acids are part of signals uses in the microbial world for cell to cell communication (Pappas et al., 2004). The FAs present in microbial lipids are mainly of four types: straight-chain saturated, straight-chain monounsaturated, branched-chain (predominantly iso and anteiso), and cyclopropane FAs. Besides these, polyunsaturated and hydroxylated FAs are abundant in some organisms (see > Fig. 1 for example of different fatty acids). In most of the naturally occurring unsaturated FAs (UFAs), the orientation about double bonds is cis rather than trans. The cis orientation has an important effect on molecular structure because each double bond inserts a bend into the hydrocarbon chain (> Fig. 1b). The nomenclature of FAs is complicated by the fact that there are at least five different systems in use. Details of FA nomenclature are explained in Gunstone and Herslo¨f (1992). The various groups of prokaryotes differ remarkably in the

Formation of Fatty Acids

26

. Figure 1 Representative structures of fatty acids.

structure and synthesis of FA-derived lipids, which serve as reliable systematic marker molecules in chemotaxonomy (Osterhout et al., 1991). Unsaturated and branched-chain (or iso-) FA increase the fluidity of the membrane and fulfill the function of thermal adaptation. The higher the content of these FAs, the lower is the solid-to-liquid phase transition temperature of these lipids.

2

Biosynthesis

Although the reaction mechanism of de novo FA biosynthesis is essentially the same in all biological systems, there are different structures of fatty acid synthase (FAS) systems. The first class is represented by the dissociated type II FAS system which occurs in most bacteria as well as in organelles (chloroplasts, mitochondria, and apicoplasts). In this case, the components are independent proteins which are encoded by a series of separated genes. In contrast, the highly integrated type I FAS multienzymes contain the various catalytic activities of the reaction sequence as discrete functional domains, either in a single polypeptide chain, or in some cases, on two different multifunctional proteins of comparable size. Type I FAS multienzymes are found characteristically in the eukaryotic cytoplasm (but also in some bacteria). Microbial type I FAS are hexamers forming either a6b6 (fungi) or a6 (bacteria) oligomers while animal

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. Figure 2 The FAS II biosynthesis pathway. See the text for details.

FAS are a2 dimers (Schweizer and Hofmann, 2004). Recently the crystallization of some of these megasynthases has been achieved (Jenni et al., 2007). In this review we will describe mainly the type II FAS (> Fig. 2). The Escherichia coli system is the paradigm for the study of this system, and X-ray structures and/or NMR structures of representative members of every enzyme of the type II pathway are now available (White et al., 2005).

2.1

The Basic Type II Fatty Acid Biosynthesis Pathway

A key feature of this system is that all fatty acyl intermediates are covalently connected to a small acidic protein, the acyl carrier protein (ACP), which shuttles them from one enzyme to another. The linkage between fatty acyl groups and ACP is achieved by a thioester bond in which the sulfhydryl of the ACP’s 40 -phosphopantetheine prosthetic group is condensed with the carboxy group of a fatty acid. Apo-ACP is converted to the active protein by holo-ACP synthase (AcpS), which transfer the 40 -phosphopantetheine from coenzyme A (CoA) to apoACP. In E. coli K12, there is a unique and essential ACP named AcpP, but other organisms have, beside the orthologous AcpP for general FA biosynthesis, other ACPs dedicated to the synthesis of specific FAs. The presence of multiple ACPs in one organism have been related to a more complex metabolism (Geiger and Lo´pez-Lara, 2002). In the FAS I system, the ACP is one

Formation of Fatty Acids

26

of the domains of the polypeptide. Fatty acid biosynthesis proceeds in two stages, initiation and cyclic elongation as outlined in > Fig. 2. For a more detailed review of the pathway and the enzymes involved, see Rock 2008 and White et al. 2005. Acetyl-CoA carboxylase (ACC) catalyzes the first committed step of FA biosynthesis. The product of the reaction is malonylCoA and the malonyl group is transferred to ACP by malonyl-CoA:ACP transacylase (FabD) to form malonyl-ACP. The condensation of malonyl-ACP with acetyl-CoA by 3-oxoacyl-ACP synthase III (FabH) with the formation of acetoacetyl-ACP and CO2 is the initiation step in FAS. Elongation of fatty acids by C2 units goes through a cycle of reactions, each cycle involving a condensation, a first reduction, a dehydratation, and a second reduction step. Each new round is initiated by an elongation condensing enzyme: FabF or FabB. FabG is the 3-oxoacyl-ACP reductase, and only a single isoform of this enzyme has been identified so far in bacteria. The next step is the dehydratation of the 3 oxoacyl-ACP to trans-2-enoyl-ACP. Two isoforms, FabA and FabZ, catalyze the dehydratation step although with different substrate specificities (Rock, 2008). Each cycle is led to completion by an enoyl-ACP reductase (FabI). Some bacteria use alternative enoyl-ACP reductases like FabL, FabK, or FabV to complete cycles of elongation (Massengo-Tiasse and Cronan, 2008; Rock, 2008).

2.2

Modifications of Fatty Acids

The 3-hydroxyacyl-ACP dehydratase FabA, not only can remove water during fatty acid elongation, but is also able to catalyze the isomerization of trans-2-decenoyl-ACP to cis-3decenoyl-ACP. For the elongation of cis-3-decenoyl-ACP to palmitoleyl-ACP FabB is essential. Bioinformatic analyses have shown that the FabA/FabB pathway for unsaturated FA might be restricted to a- and g-proteobacteria (Rock, 2008). In Streptococci, after the formation of trans-2-decenoyl-ACP by FabZ, the specific isomerase FabM isomerizes it to cis-3-decenoylACP. Surprisingly, in Enterococcus FabZ has isomerase activity and FabZ/FabF of Enterococcus can functionally replace the FabA/FabB system of E. coli (Wang and Cronan, 2004). Another pathway for unsaturated FA synthesis in bacteria occurs by aerobic desaturation. In this case, the fatty acid is modified postbiosynthetically when fatty acyl residues are already attached to membrane phospholipids. This system is found in bacilli and cyanobacteria and is a mechanism conserved in Bacteria and Eukarya (Aguilar and de Mendoza, 2006). Not all bacteria regulate membrane fluidity through the production of monounsaturated straight-chain FAs. Most Gram-positive bacteria use branched-chain FAs to modulate membrane fluidity. The branch is a methyl group in the iso or anteiso-position in the chain (i.e., on the second or third carbon from the distal end of the chain, > Fig. 1d). The branched chains are introduced into the pathway in the initiation phase by FabH enzymes with different specificity to that of E. coli. FabH of E. coli is selective for acetyl-CoA while each of the FabH isoenzymes of Bacillus subtillis prefers the branched-chain substrates isobutyryl-CoA or 2-methylvaleryl-CoA (Rock, 2008). FabH from Mycobacterium tuberculosis has the largest acyl-chain-binding pocket, and can accommodate acyl-CoAs of 14 carbons and longer (Choi et al., 2000). The presence of 3-hydroxylated FAs is common in some bacteria and these fatty acids are the primary fatty acids in Lipid A (Raetz and Whitfield, 2002) as well as in ornithinecontaining lipids (OL) (Lo´pez-Lara et al., 2003). 3-hydroxy-fatty acyl-ACPs are normal intermediates in the FAS II elongation cycle (> Fig. 2). The acyltransferases involved in the addition of the primary fatty acids in lipid A biosynthesis (LpxA and LpxD) or in OL

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biosynthesis (OlsB) must have a higher affinity for the specific 3-hydroxy-fatty acyl-ACP than the FASII enzymes and thus efficiently remove the intermediate from the biosynthesis pathway. In contrast, S-2-hydroxylation usually is introduced after the fatty acyl group has been attached to the lipid A molecule and is catalyzed by the LpxO dioxygenase (Gibbons et al., 2008). Similar dioxygenases might be responsible for the introduction of 2-hydroxy substitutions on the fatty acyl group of OL, bacterial sphingolipids, or phosphatidyletanolamine (Geiger et al., 2009). In numerous representatives of the a-2 subgroup of the proteobacteria, the lipid A carries a long (omega-1)-hydroxy fatty acid named 27-hydroxyoctacosanoic acid (C28). Interestingly, a specialized ACP, AcpXL, is required for the transfer of C28 to lipid A by the acyltransferase LpxXL (Basu et al., 2002). Between the genes for acpXL and lpxXL, four genes are located that encode homologues of fatty acid biosynthetic enzymes (Basu et al., 2002) and therefore presumably there is a complete set of FASII enzymes dedicated to the biosynthesis of C28 where the acyl intermediates will be bound to the specialized AcpXL, instead than to the housekeeping AcpP. The conversion of preexisting cis-unsaturated FAs to cyclopropane fatty acids (i.e., lactobacillic acid shown in > Fig. 1e) is widespread in bacteria. The required methylation is carried out by cyclopropane fatty acid synthase (Cfa), which uses S-adenosylmethionine as the methyl donor to create the cyclopropane ring (Rock, 2008). A few bacteria have evolved a mechanism to convert existing cis-unsaturated FAs in phospholipids into trans-UFAs to adapt to environmental challenges. The isomerization is carried out by periplasmic cis-trans isomerase (Cti) (Holtwick et al., 1997). For the synthesis of the unusual a-b unsaturated fatty acids, which are specific substituents of nodulation (Nod) factors (> Fig. 3e), it seems that a specialized ACP, NodF, and a different elongation condensing enzyme, NodE, work in combination with enzymes of the basic FAS II system (Geiger and Lo´pez-Lara, 2002).

3

Fatty Acids and Fatty Acid-Derivatives as Signal Molecules

In some cases, fatty acids themselves or more commonly fatty acid-containing molecules are functioning in bacteria as chemical signals for cell-to-cell communication, a process that is known as quorum sensing. These signals include 3-hydroxypalmitic methyl ester (isolated from Ralstonia solanacearum), cis-11-methyl-2-dodecenoic acid (named the diffusible signal factor (DSF) which is probably also involved in interspecies signalling (Ryan et al., 2008)), acylated homoserine lactones (AHLs) in the Proteobacteria and g-butyrolactones of filamentous Streptomyces (> Fig. 3). Also the molecules that rhizobia use to trigger the development of nodules on the roots of their respective host-plant are acylated oligosaccharides (> Fig. 3e). Furthermore, in acylated signals such us AHLs and Nod factors, the fatty acyl chains are important determinants of the specificity (i.e., Demont-Caulet et al., 1999). AHL synthases (LuxI-like enzymes) produced AHLs from the substrates S-adenosyl-L-methionine (SAM) and acyl-ACP and the choice of fatty acyl substrate is due to the substrate preference of the enzyme (reviewed in Pappas et al., 2004). A-factor (2-isocapryloyl-3R-hydroxymethyl-gbutyrolactone, > Fig. 3d) is a representative of the g-butyrolactone autoregulators that trigger secondary metabolism and morphogenesis in Streptomyces. The biosynthesis pathway for A-factor has been reported and an acyl-ACP is one of the precursors (Kato et al., 2007).

Formation of Fatty Acids

26

. Figure 3 Structures of two fatty acids that function as signals molecules (a and b) and representatives of families of acyl-based signalling molecules of the AHLs (c), g-butyrolactones (d) and of Nod factors (e).

4

Specificity of Acyltransferases

In the biosynthesis of phospholipids, the bacterial glycerol-phosphate acyltransferases utilize the completed fatty acyl chains to form membrane phospholipids. Initially, the bacterial acyltransferases PlsB and PlsC of E. coli were studied and it was shown that these two acyltransferases use thioesters of either CoA or acyl-ACP as the acyl donors (Rock, 2008). However, most organisms use the recently discovered PlsX/PlsY system, instead of the PlsB enzyme. Although PlsC is widely distributed, the PlsC homologue in Bacillus subtilis can use only acyl-ACP (Paoletti et al., 2007). LpxA is the acyltransferase that catalyzes the first step of lipid A biosynthesis, the reversible transfer of the R-3-hydroxyacyl chain from R-3hydroxyacyl-ACP to the glucosamine 3-OH group of UDP-GlcNAc. E. coli LpxA is highly selective for R-3-hydroxymyristate. Recently the structural basis of LpxA specificity for the

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Formation of Fatty Acids

fatty acid was demonstrated (Williams and Raetz, 2007). AHLs vary greatly in acyl-chain length, from C4-AHL up to C18-AHLs and each AHL synthase produces specifically a type of AHL. Sequence analysis of the AHL-synthase family fails to reveal a robust correlation between sequence composition and acyl-chain length. Therefore, to accommodate and create a preference for acyl-ACPs of different acyl-chain lengths, a number of sequence and tertiary structure differences must have evolved in different AHL synthases (Gould et al., 2004). In the AHL EsaI that produces 3-oxo-C6-AHL a restrictive hydrophobic pocket was found (Watson et al., 2002) in contrast to the presence of a more relaxed hydrophobic tunnel present in the AHL LasI that produces 3-oxo-C12-AHL (Gould et al., 2004).

5

Research Needs

FAS II has been extensively studied in the E. coli model system and with the availability of hundreds of genome sequences, the pathway has been investigated in other organisms, mostly important human pathogens. In this way, it will be interesting to know the similarities and differences with respect to the E. coli pathway, not only from the point of view of basic research, but also because FAS II is emerging as a major target for the development of novel antibacterial agents (Zhang et al., 2006). The focus is on the development of broad spectrum inhibitors targeting bacterial FAS II and not the structurally different mammalian type I FAS. On the other hand, efforts are needed to find specific inhibitors of FAS in the important pathogens Mycobacterium tuberculosis and malaria (reviewed in Zhang et al., 2006). The structures of the different components of the FAS II systems of E. coli are known and these data will be very helpful to identify the mechanisms of interaction of ACP with the different enzymes. Rather than an ACP-binding motif defined by primary structure, the enzymes of the type II system share 3D surface features that account for their specific recognition of ACP and its thioesters (Zhang et al., 2003). Also, a putative binding site for ACP on the surface of the AHL synthase LasI includes a basic patch of residues (Gould et al., 2004). Cocrystallization of the different enzymes with their respective acyl-ACP will help to confirm the structure of the ACP binding site as well as to determine in more examples which structural features determine the specificity for the acyl chain. Finally, the structural basis of the interaction of specialized ACPs with their corresponding enzymes and acyl chains should be investigated in order to understand the contribution of these specialized ACPs in channeling biosynthetic building blocks into certain pathways.

References Aguilar PS, de Mendoza D (2006) Control of fatty acid desaturation: A mechanism conserved from bacteria to humans. Mol Microbiol 62: 1507–1514. Alvarez H, Steinbu¨chel MA (2002) Triacylglycerols in prokaryotic microorganisms. Appl Microbiol Biot 60: 367–376. Basu SS, Karbarz MJ, Raetz CR (2002) Expression cloning and characterization of the C28 acyltransferase of lipid A biosynthesis in Rhizobium leguminosarum. J Biol Chem 277: 28959–28971.

Choi KH, Kremer L, Besra GS, Rock CO (2000) Identification and substrate specificity of beta-ketoacyl (acyl carrier protein) synthase III (mtFabH) from Mycobacterium tuberculosis. J Biol Chem 275: 28201–28207. Demont-Caulet N, Maillet F, Tailler D, Jacquinet JC, Prome´ JC, Nicolaou KC, Truchet G, Beau JM, De´narie´ J (1999) Nodule-inducing activity of synthetic Sinorhizobium meliloti nodulation factors and related lipo-chitooligosaccharides on alfalfa.

Formation of Fatty Acids Importance of the acyl chain structure. Plant Physiol 120: 83–92. Geiger O, Lo´pez-Lara IM (2002) Rhizobial acyl carrier proteins and their roles in the formation of bacterial cell-surface components that are required for the development of nitrogen-fixing root nodules on legume hosts. FEMS Microbiol Lett 208: 153–162. Geiger O, Sohlenkamp C, Lo´pez-Lara IM (2009) Formation of bacterial membrane lipids: pathways, enzymes, reactions. This book, Chapter 70. Gibbons HS, Reynolds CM, Guan Z, Raetz CR (2008) An inner membrane dioxygenase that generates the 2-hydroxymyristate moiety of Salmonella lipid A. Biochemistry 47: 2814–2825. Gould TA, Schweizer HP, Churchill ME (2004) Structure of the Pseudomonas aeruginosa acylhomoserinelactone synthase LasI. Mol Microbiol 53: 1135–1146. Gunstone FD, Herslo¨f BG (1992) A lipid glossary. Great Britain : The Oily Press. Holtwick R, Meinhardt F, Keweloh H (1997) Cis-trans isomerization of unsaturated fatty acids: Cloning and sequencing of the cti gene from Pseudomonas putida P8. Appl Environ Microbiol 63: 4292–4297. Jenni S, Leibundgut M, Boehringer D, Frick C, Mikola´sek B, Ban N (2007) Structure of fungal fatty acid synthase and implications for iterative substrate shuttling. Science 316: 254–261. Kato JY, Funa N, Watanabe H, Ohnishi Y, Horinouchi S (2007) Biosynthesis of gamma-butyrolactone autoregulators that switch on secondary metabolism and morphological development in Streptomyces. Proc Natl Acad Sci USA 104: 2378–2383. Lo´pez-Lara IM, Sohlenkamp C, Geiger O (2003) Membrane lipids in plant-associated bacteria: their biosyntheses and possible functions. Mol Plant Microbe Interact 16: 567–579. Massengo-Tiasse´ RP, Cronan JE (2008) Vibrio cholerae FabV defines a new class of enoyl-acyl carrier protein reductase. J Biol Chem 283: 1308–1316. Neuhaus FC, Baddiley J (2003) A continuum of anionic charge: Structures and functions of D-alanylteichoic acids in gram-positive bacteria. Microbiol Mol Biol Rev 67: 686–723. Osterhout GJ, Shull VH, Dick JD (1991) Identification of clinical isolates of gram-negative nonfermentative bacteria by an automated cellular fatty acid identification system. J Clin Microbiol 29: 1822–1830. Paoletti L, Lu YJ, Schujman GE, de Mendoza D, Rock CO (2007) Coupling of fatty acid and phospholipid synthesis in Bacillus subtilis. J Bacteriol 189: 5816–5824. Pappas KM, Weingart CL, Winans SC (2004) Chemical communication in proteobacteria: Biochemical and structural studies of signal synthases and receptors

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required for intercellular signalling. Mol Microbiol 53: 755–769. Pugh EL, Kates M (1994) Acylation of proteins of the archaebacteria Halobacterium cutirubrum and Methanobacterium thermoautotrophicum. Biochim Biophys Acta 1196: 38–44. Raetz CR, Reynolds CM, Trent MS, Bishop RE (2007) Lipid A modification systems in Gram-negative bacteria. Annu Rev Biochem 76: 295–329. Raetz CR, Whitfield C (2002) Lipopolysaccharide endotoxins. Annu Rev Biochem 71: 635–700. Rezwan M, Grau T, Tschumi A, Sander P (2007) Lipoprotein synthesis in mycobacteria. Microbiology 153: 652–658. Ryan RP, Fouhy Y, Garcia BF, Watt SA, Niehaus K, Yang L, Tolker-Nielsen T, Dow JM (2008) Interspecies signalling via the Stenotrophomonas maltophilia diffusible signal factor influences biofilm formation and polymyxin tolerance in Pseudomonas aeruginosa. Mol Microbiol 68: 75–86. Rock CO (2008) Fatty acids and phospholipids metabolism in prokaryotes. In Biochemistry of lipids, lipoproteins, and membranes, 5th edn. DE Vance, JE Vance (eds.). Amsterdam: Elsevier, pp 59–96. Schweizer E, Hofmann J (2004) Microbial type I fatty acid synthases (FAS): Major players in a network of cellular FAS systems. Microbiol Mol Biol R 68: 501–517. Steinbu¨chel A (2001) Perspectives for biotechnological production and utilization of biopolymers: Metabolic engineering of polyhydroxyalkanoate biosynthesis pathways as a successful example. Macromol Biosci 1: 1–24. Wang H, Cronan JE (2004) Functional replacement of the FabA and FabB proteins of Escherichia coli fatty acid synthesis by Enterococcus faecalis FabZ and FabF homologues. J Biol Chem 279: 34489–33495. Watson WT, Minogue TD, Val DL, Beck von Bodman S, Churchill MEA (2002) Structural basis and specificity of acyl-homoserine lactone signal production in bacterial quorum sensing. Mol Cell 9: 685–694. White SW, Zheng J, Zhang YM, Rock CO (2005) The structural biology of type II fatty acid biosynthesis. Annu Rev Biochem 74: 791–831. Williams AH, Raetz CR (2007) Structural basis for the acyl chain selectivity and mechanism of UDP-Nacetylglucosamine acyltransferase. P Natl Acad Sci USA 104: 13543–13550. Zhang YM, Marrakchi H, White SW, Rock CO (2003) The application of computational methods to explore the diversity and structure of bacterial fatty acid synthase. J Lipid Res 44: 1–10. Zhang YM, White SW, Rock CO (2006) Inhibiting bacterial fatty acid synthesis. J Biol Chem 281: 17541–17544.

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27 Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions O. Geiger* . C. Sohlenkamp . I. M. Lo´pez-Lara Centro de Ciencias Geno´micas, Universidad Nacional Auto´noma de Me´xico, Morelos, Me´xico *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 396

2

Membrane-Forming Lipids in Escherichia coli . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397

3 3.1 3.2 3.2.1 3.2.2 3.2.3 3.2.4 3.2.5 3.2.6

Formation of Glycerophospholipids in Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 397 Formation of the Central Activated Intermediate CDP-Diacylglycerol . . . . . . . . . . . 397 Diversification of Polar Head Groups in Bacterial Glycerophospholipids . . . . . . . . 399 Biosynthesis of Phosphatidylglycerol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 Biosynthesis of Lysyl-Phosphatidylglycerol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 Biosynthesis of Cardiolipin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 Biosynthesis of Phosphatidylethanolamine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 399 Biosynthesis of Phosphatidylcholine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400 Biosynthesis of Phosphatidylinositol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400

4

Formation of Sphingolipids in Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 400

5

Steroid and Hopanoid Formation in Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 401

6 6.1 6.2 6.3 6.4

Formation of Phosphorus-Free Membrane Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402 Glycolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 402 Sulfolipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403 Betaine Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403 Ornithine-Containing Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404

7

Phenolic Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 404

8

Conclusions and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 405

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_27, # Springer-Verlag Berlin Heidelberg, 2010

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Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

Abstract: The model bacterium Escherichia coli contains the phospholipids phosphatidylglycerol, cardiolipin, and phosphatidylethanolamine as major membrane lipids and biosyntheses and functionalities of individual membrane lipids have mainly been studied in this organism. However, in other bacteria, additional and alternative membrane lipids are found and in many cases neither their biosyntheses nor their functionalities are understood. Some Gram-negative bacteria have phosphatidylcholine or sphingolipids in their standard repertoire, whereas many Gram-positives have glycosylated diacylglycerols and lysyl-phosphatidylglycerol in their membranes. Notably, phosphatidylinositol is an essential lipid for Mycobacterium tuberculosis. Steroid and hopanoid lipids only occur in some bacteria. Under certain stress conditions specific membrane lipids can be formed in order to minimize the stress exerted. For example, under phosphorus-limiting conditions of growth, some bacteria form membrane lipids lacking phosphorus such as glycolipids, sulfolipids, betaine lipids, or ornithine-containing lipids. Challenge of proteobacteria with acid causes modifications of membrane lipids, such as formation of lysyl-phosphatidylglycerol or hydroxylation of ornithine-containing lipids.

1

Introduction

A primary role of lipids in cellular function is to form the lipid bilayer permeability barrier of cells (See > Chapter 29, Vo1. 1, Part 5). Also, there are at least two ways by which lipids can affect protein structure and function and thereby function of the cell. In one way, protein function is influenced by specific protein-lipid interactions that depend on the chemical and structural anatomy of lipids (head group, backbone, alkyl chain length, degree of unsaturation, chirality, ionization, and chelating properties). In another way, protein function is also influenced by the unique self-association properties of lipids that result from the collective properties (fluidity, bilayer thickness, shape, and packing properties) of the lipids organized into membrane structures. Glycerophospholipids are the primary building blocks of membranes but other lipids are important components. Eukaryotic membranes contain phospholipids (glycerophospholipids and phospho-sphingolipids), phosphorus-free glycerolipids and sphingosin-based lipids as well as steroids (Nelson and Cox, 2008). Although highly decorated sphingolipid derivatives or the broad variety of steroids are absent in prokaryotes, Bacteria possess all the basic structures of membrane lipids found in Eukarya, albeit not in one single species. In Eukarya, sphingolipids and cholesterol cluster together in membrane lipid rafts (See > Chapter 62, Vol. 4, Part 7) and numerous specific functions are associated with lipid rafts (Nelson and Cox, 2008). Defined lipid domains exist also in Bacteria (Matsumoto et al., 2006) and recently it has been shown that the cell division-inhibiting protein MinD is associated with phosphatidylglycerol-containing lipid spirals in Bacillus (Bara´k et al., 2008). The membrane lipids of Archaea (See > Chapter 33, Vol. 1, Part 5) differ from those of Bacteria and Eukarya. In contrast to the lipids of Bacteria and Eukarya in which ester linkages bond the fatty acids to glycerol, the lipids of Archaea contain ether bonds between glycerol and their hydrophobic side chains. In addition, archaeal lipids lack fatty acids. Instead, the side chains are composed of repeating units of isoprene (Dowhan et al., 2008). In this review we will focus on membrane-forming lipids of Bacteria. Gram-negative bacteria possess functionally distinct inner (cytoplasmic) and outer membranes (Dowhan

Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

27

et al., 2008). However, recent evidence suggests that in addition to their cytoplasmic membrane also some Gram-positive bacteria might have an outer membrane bilayer structure formed by long-chain mycolic acids and other membrane lipids (Hoffmann et al., 2008).

2

Membrane-Forming Lipids in Escherichia coli

In the bacterial model organism, Escherichia coli, only the three major membrane lipids phosphatidylethanolamine, phosphatidylglycerol, and cardiolipin occur (See > Chapter 28, Vol. 1, Part 5). In addition, Gram-negative bacteria usually have the lipid A-containing lipopolysaccharide in the outer monolayer of their outer membrane and lipid A modification systems have been reviewed recently (Raetz et al., 2007). However, even in E. coli many minor membrane phospholipids exist (Raetz, 1986). For many of these minor phospholipids, neither their structures nor the genes/enzymes required for their biosyntheses are known.

3

Formation of Glycerophospholipids in Bacteria

3.1

Formation of the Central Activated Intermediate CDP-Diacylglycerol

In Bacteria, glycerol-3-phosphate (> Fig. 1) forms the backbone of all glycerophospholipid molecules. It can be synthesized by two different pathways, from glycerol directly or from the glycolytic intermediate dihydroxyacetone phosphate (DHAP). During growth with glycerol as sole carbon source, the enzymes of the glycerol catabolic (glp) operon are induced. One of the induced enzymes, a glycerol kinase (GlpK), is able to phosphorylate glycerol and form glycerol-3-phosphate. During growth on carbon sources other than glycerol, glycerol-3-phosphate is made by direct reduction of DHAP with NADH. The enzyme catalyzing this reaction is called biosynthetic glycerol-3-phosphate dehydrogenase (GpsA) (Rock, 2008). In E. coli, the glycerol-3-phosphate acyltransferase PlsB can use acyl-CoA or acyl-ACP (See > Chapter 26, Vol. 1, Part 5) as acyl donors and is the major activity for catalyzing the first acylation at position 1 of glycerol-3-phosphate thereby forming 1-acyl-glycerol-3-phosphate. However, the more widespread pathway to achieve the initial acylation of glycerol-3phosphate among bacteria seems to involve PlsX and PlsY (Lu et al., 2006). In this novel pathway, PlsX catalyzes the conversion of acyl-ACP and inorganic phosphate to acylphosphate and ACP. In a second step, PlsY transfers the acyl group from acyl-phosphate to glycerol-3-phosphate forming inorganic phosphate and 1-acyl-glycerol-3-phosphate (Lu et al., 2006). The second fatty acyl residue is added by another enzyme, the 1-acyl-glycerol-3-phosphate acyltransferase PlsC, to form phosphatidic acid. Although PlsC from E. coli can use acylACP or acyl-CoA as acyl donors, PlsC from many other bacteria might use exclusively acyl-ACP (Lu et al., 2006). There is a tendency that most fatty acids at the 1-position are saturated whereas most at the 2-position are unsaturated. The acylation specificity, however, is not absolute and can be altered by the supply of acyl donors. The conversion of phosphatidic acid to CDP-diacylglycerol (CDP-diglyceride) is catalyzed by CDP-diglyceride synthase CdsA (Rock, 2008) (> Fig. 1).

397

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Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

. Figure 1 Model of glycerophospholipid biosynthesis in bacteria (for details see text).

Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

3.2

Diversification of Polar Head Groups in Bacterial Glycerophospholipids

3.2.1

Biosynthesis of Phosphatidylglycerol

27

Phosphatidylglycerol phosphate synthase (PgsA) transfers glycerol-3-phosphate to CDPdiacylglycerol under the release of CMP thereby producing phosphatidylglycerol phosphate (PGP). There are at least two enzymes with PGP phosphatase activity (PgpA and PgpB) in E. coli, releasing inorganic phosphate from PGP to form phosphatidylglycerol (PG) (Rock, 2008) (> Fig. 1). PG or other negatively charged membrane lipids are required for initiation of DNA replication, control of cell division, and protein transloction across membranes (Dowhan et al., 2008).

3.2.2

Biosynthesis of Lysyl-Phosphatidylglycerol

Lysyl-phosphatidylglycerol (lysyl-PG) is a well-known membrane lipid in many Gram-positive bacteria (Staphylococcus, Lactococcus, Bacillus) and its formation increases the resistance of bacteria against cationic peptides of the innate immune response. The staphylococcal MprF can transfer lysine from charged lysyl-tRNA to PG forming lysyl-PG. In some Gram-negative bacteria, the MprF homologue LpiA is induced under acidic conditions forming lysyl-PG which seems to increase the resistance to cationic peptides (Sohlenkamp et al., 2007)(> Fig. 1). The pathogen Clostridium perfringens has two phylogenetically distinct MprF paralogs, one responsible for the formation of lysyl-PG and the other causing the synthesis of alanylphosphatidylglycerol (Roy and Ibba, 2008).

3.2.3

Biosynthesis of Cardiolipin

In E. coli and probably most other bacteria, a cardiolipin synthase (ClsB) of the prokaryotic type condenses two PG molecules to yield cardiolipin (CL) and free glycerol in a transesterification reaction (> Fig. 1). However, inactivation of a mycobacterial gene coding for a presumptive CDP-alcohol phosphatidyltransferase led to a reduced formation of cardiolipin (Jackson et al., 2000), provoking the suggestion that some bacteria might use a eukaryotic type cardiolipin synthase (ClsE) which condenses CDP-diacylglycerol with PG to form cardiolipin and CMP. Recent evidence suggests that cardiolipin controls the osmotic stress response and the subcellular location of the transporter ProP in E. coli (Romantsov et al., 2008).

3.2.4

Biosynthesis of Phosphatidylethanolamine

The first step in the synthesis of phosphatidylethanolamine (PE) is the condensation of CDPdiacylglycerol with serine to form phosphatidylserine (PS) catalyzed by PS synthase (Pss). In a second step, the decarboxylation of PS is catalyzed by PS decarboxylase (Psd) to yield PE (> Fig. 1). PE is required in E. coli for the proper folding and topological organization of certain membrane proteins within the lipid bilayer. For example, permeases, such as the

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Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

lactose permease LacY, require the presence of PE for active transport of their substrates (Dowhan et al., 2008).

3.2.5

Biosynthesis of Phosphatidylcholine

Although phosphatidylcholine (PC; lecithin) is the major membrane-forming phospholipid in Eukarya, it is thought to be present in only about 10% of the bacteria (Sohlenkamp et al., 2003). A well-known pathway for PC formation occurs by three-fold methylation of PE using S-adenosylmethionine (SAM) as methyl donor and catalyzed by phospholipid N-methyltransferase (PmtA). In many bacteria, all three subsequent methylations are achieved by a single enzyme (> Fig. 1). Some complex bacteria with large genomes, however, such as Bradyrhizobium japonicum, possess up to four distinct phospholipid N-methyltransferases with distinct substrate specificities. B. japonicum has phospholipid N-methyltransferases that predominantly perform the first, the second, or the third methylation (Hacker et al., 2008). Many PCcontaining bacteria have a second pathway for PC formation, catalyzed by PC synthase (Pcs), in which choline is condensed directly to CDP-diacylglycerol forming PC and CMP (Sohlenkamp et al., 2003) (> Fig. 1). To date, Pcs has been found only in bacteria. Eukarya usually have another pathway, the CDP-choline pathway to recycle free choline to PC. In the CDPcholine pathway, choline is phosphorylated to choline phosphate by choline kinase and further activated by CTP:phosphocholine cytidylyltransferase to CDP-choline. In a third step, catalyzed by CDP-choline:1,2-diacylglycerol choline phosphotransferase, the CDP-choline is condensed to the non-activated lipid part 1,2-diacylglycerol forming PC and CMP. This latter eukaryote-like pathway might exist in some bacteria such as Treponema (Kent et al., 2004). In Legionella pneumophila (See > Chapter 64, Vol. 4, Part 7), bacterial PC seems to be required for multiple functions, such as attachment to the macrophage via the platelet-activating factor receptor, timely formation of a functioning type IV secretion system, as well as transition of the bacterium to a state of higher virulence (Conover et al., 2008).

3.2.6

Biosynthesis of Phosphatidylinositol

Phosphatidylinositol (PI), a membrane lipid central to signaling in eukaryotes, is absent in most bacteria. However, the causative agent of tuberculosis Mycobacterium tuberculosis has PI and derivatives thereof as major components in its membrane. In M. tuberculosis, PI is formed by condensing myo-inositol to CDP-diacylglyceride in a reaction catalyzed by PI synthase (Jackson et al., 2000) (> Fig. 1). PI formation by PI synthase is essential for the survival of Mycobacterium and therefore the mycobacterial PI synthase might be a promising drug target for a antituberculosis therapy (Jackson et al., 2000). Within the Bacteria, PI synthase and therefore PI seems to be limited to actinobacteria (Michell, 2008).

4

Formation of Sphingolipids in Bacteria

In eukaryotes, sphingolipids are crucial for signaling and organization of lipid rafts. Many Bacteroides species have two types of phosphosphingolipids, ceramide phosphorylethanolamine

Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

27

and ceramide phosphorylglycerol (Kato et al., 1995). However, in bacteria the occurrence of sphingolipids is so unusual that if they are detected, it is thought so remarkable that often the genus name of the respective bacterium harbours the prefix ‘‘Sphingo,’’ i.e., in Sphingomonas and Sphingobacterium. Some Gram-negative bacteria which lack lipopolysaccharides in their outer membranes have glycosylated sphingolipids as functional replacements. These glycosphingolipids of the outer bacterial membrane are recognized by natural killer T cells which provide an innate-type immune response towards glycosphingolipid-containing bacteria (Wu et al., 2006). So far only the first step in bacterial sphingolipid biosynthesis is well characterized. The transacylation from palmitoyl-CoA to serine yielding CoA, CO2, and 3-ketodihydrosphingosine is catalyzed by serine palmitoyltransferase of which even the structure has been resolved (Yard et al., 2007).

5

Steroid and Hopanoid Formation in Bacteria

While sterol synthesis is nearly ubiquitous among eukaryotes, only very few reports on the presence of steroids in phylogenetically distant bacterial taxa exist (Summons et al., 2006) (See > Chapter 7, Vol. 1, Part 2). The key enzyme oxidosqualene cyclase (Osc) cyclizes squalene (3S)-2,3-epoxide to the tetracyclic core that characterizes all sterols (Summons et al., 2006) (> Fig. 2). Hopanoids (See > Chapter 8, Vol. 1, Part 2) are pentacyclic triterpenoid lipids occurring predominantly in bacteria. They have been shown to influence membrane ordering and fluid

. Figure 2 Biosynthesis of steroids and hopanoids in bacteria (for details see text).

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Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

properties similarly to steroids and therefore hopanoids have been termed ‘‘sterol surrogates’’ suggesting that they might be functionally equivalent to sterols in membranes. Although steroids and hopanoids are structurally similar, they differ in a key respect: steroids have their polar group attached directly to the ring structure at C3, while the polar functions in hopanoids are attached to the side chain which must have distinct implications for their movements within membranes (Summons et al., 2006) (> Fig. 2). Although hopanoids are widely distributed in bacteria, little is known about their formation and their molecular functions. The key enzyme squalene-hopene cyclase (SqhC) converts the isoprenoid-derived squalene into the cyclic form of hopene (> Fig. 2). So far the role of hopanoids in nitrogenfixing organisms is unclear. In some rhizobial bacteria, such as Bradyrhizobium japonicum which forms nitrogen-fixing root nodules with soja host plants, hopanoids can make up to 50% (w/w) of the total lipid fraction. Surprisingly however, no triterpenoids have been detected in the fast-growing genera Rhizobium or Sinorhizobium and these genera seem to lack a gene for SqhC. In the Gram-positive bacterium Frankia, nitrogen fixation occurs within specialized multicellular structures termed vesicles. A vesicle is surrounded by a multilamellate, lipid-containing envelope that apparently functions as a barrier to oxygen diffusion. The envelope consists primarily of two hopanoid lipids, bacteriohopanetetrol and bacteriohopanetetrol phenylacetate monoester, the latter of which is vesicle-specific (Berry et al., 1993). Hopanoids are also formed during transition from substrate to aerial hyphae in Streptomyces (Poralla et al., 2000).

6

Formation of Phosphorus-Free Membrane Lipids

The composition of lipids in the cell membrane can vary depending on the physiological situation of an organism (Lo´pez-Lara et al., 2003). Under phosphorus-limiting conditions, membrane phospholipids of some bacteria are at least partially replaced by lipids containing no phosphorus as demonstrated in Bacillus subtilis, Pseudomonas diminuta, Pseudomonas fluorescens, and Rhodobacter sphaeroides (reviewed in Lo´pez-Lara et al., 2003). In Sinorhizobium (Rhizobium) meliloti these phosphorus-free lipids are the sulfolipid sulfoquinovosyl diacylglycerol (SL), ornithine-containing lipid (OL), and the betaine lipid diacylglyceryl trimethylhomoserine (DGTS) and in some other bacteria these phosphorus-free lipids include glycolipids. Notably, glycolipids, sulfolipids, and betaine lipids are all formed from the same precursor diacylglycerol (> Fig. 3).

6.1

Glycolipids

Glycosyl diacylglycerols are widespread membrane glycolipids in plants, animals and Grampositive bacteria but they have not been frequently reported in Gram-negative bacteria. The ypfP-encoded UDP glucosyltransferase from Bacillus subtilis successively transfers up to four glucose residues from UDP-glucose to diacylglycerol (Jorasch et al., 1998) (> Fig. 3). An essential glycosyltransferase (MPN483) from the human pathogen Mycoplasma pneumoniae is also processive and relatively flexible with regard to its substrates. MPN483 can use UDPgalactose or UDP-glucose as sugar donors and diacylglycerol or ceramide derivatives as acceptors thereby producing a wide spectrum of compounds, some of them important antigens during infection by Mycoplasma (Klement et al., 2007).

Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

27

. Figure 3 Biosynthesis of diacylglycerol-derived phosphorus-free membrane lipids in bacteria (for details see text).

6.2

Sulfolipids

The sulfolipid sulfoquinovosyl diacylglycerol occurs widely in photosynthetic organisms ranging from bacteria to seed plants (Benning, 1998) where it is associated with photosynthetic membranes. Although sulfolipid is not generally essential for photosynthesis, it is required for growth under phosphate-limiting growth conditions in R. sphaeroides and cyanobacteria. In R. sphaeroides, phosphate limitation of the wild type caused a significant reduction in the amount of all phospholipids and an increased amount of sulfolipid. This sulfolipid, 6-sulfo-a-D-quinovosyl diacylglycerol, has been suggested to function as a surrogate for phospholipids, particularly phosphatidylglycerol, under phosphate-limiting conditions (Benning et al., 1993). At least 4 structural genes (sqdA, sqdB, sqdC, and sqdD) are involved in sulfolipid biosynthesis in R. sphaeroides (Benning, 1998) (> Fig. 3). The gene sqdB encodes for an enzyme involved in the biosynthesis of UDP-sulfoquinovose from UDPglucose and sulfite as the sulfur donor. Although sqdA is predicted to encode an acyltransferase its precise biochemical function remains unclear (Benning, 2007). Finally, products of sqdC and sqdD are thought to catalyze the transfer of sulfoquinovose from UDP-sulfoquinovose to diacylglycerol but the detailed overall mechanism for sulfolipid assembly in bacteria has not been resolved (Benning, 2007).

6.3

Betaine Lipids

Although PC is known to be the major membrane lipid in eukaryotes, some lower eukaryotic organisms possess the betaine lipid diacylglyceryl-N,N,N-trimethylhomoserine (DGTS)

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instead. DGTS occurrs in a wide variety of lower green plants (green algae, bryophytes and pteridophytes), chromophytes, fungi, and amoebae (reviewed in Sohlenkamp et al., 2003). In some a-proteobacteria, DGTS has been identified as a phosphorus-free membrane lipid that substitutes for PC under conditions of phosphate limitation. There is an apparent reciprocity between the content of PC and the content of DGTS, i.e., when PC is a major membrane lipid often no DGTS is detected in the same organism, whereas in organisms where DGTS is a major lipid, PC is only found in trace levels. This suggests, that DGTS and PC, both zwitterionic at physiological pH, are interchangeable at least with regard to essential functions for the respective organism. Two structural genes from Rhodobacter sphaeroides, btaAB, coding for two enzymes BtaA and BtaB involved in DGTS biosynthesis have been characterized (Riekhof et al., 2005). The BtaA S-adenosylmethionine/diacylglycerol 3-amino-3-carboxypropyl transferase converts diacylglycerol (DAG) into diacylglyceryl-homoserine (DGHS) and during the formation of the ether bond, S-adenosylmethionine functions as donor of the homoseryl group. Subsequently, the S-adenosylmethionine:diacylglycerylhomoserine-N-methyltransferase BtaB catalyzes threefold methylation of DGHS in order to yield DGTS (> Fig. 3).

6.4

Ornithine-Containing Lipids

Ornithine-containing lipids (OL) are widespread among Gram-negative bacteria and occur in some Gram-positives, like Mycobacterium and Streptomyces species (reviewed in Lo´pez-Lara et al., 2003) but are absent in Archaea and Eukarya. The structures of OLs have been reported from a number of organisms and were found to be a-N-(acyloxyacyl)-ornithines. In this structure, a 3-hydroxyfatty acyl group is attached in amide linkage to the a-amino group of ornithine. The synthesis of OL occurs in two steps. In a first step, the N-acyltransferase OlsB transfers 3-hydroxy-fatty acyl residue from 3-hydroxy-fatty acyl-ACP to ornithine in order to obtain lyso-ornithine lipid (Gao et al., 2004). In a second step, lyso-ornithine-lipid is converted to OL by an olsA-dependent O-acyltransferase activity that requires acyl-AcpP as the acyl donor. In some bacteria, like Burkholderia cepacia, Flavobacterium, or Rhizobium tropici besides the OL mentioned above, OL exist in which the ester-linked fatty acyl group possesses a hydroxyl group at the 2-position (Rojas-Jime´nez et al., 2005). Similar S-2-hydoxyfatty acyl moieties are integral parts of Salmonella typhimurium lipid A and are thought to be of importance for pathogenesis of this organism. The S-2-hydroxylation is introduced after the fatty acyl group had been attached to the lipid A molecule and is catalyzed by the LpxO dioxygenase (Gibbons et al., 2008). A similar dioxygenase (OlsC) might be responsible for the introduction of 2-hydroxy substitutions on the ester-linked fatty acyl group of OL under conditions of acid stress (Rojas-Jime´nez et al., 2005). Notably, 2-hydroxy substitutions on ester-linked fatty acyl groups have also been reported for bacterial sphingolipids and PE, other major components of the outer membrane in Gram-negative bacteria > Fig. 4.

7

Phenolic Lipids

Phenolic lipids, (See > Chapter 32, Vol. 1, Part 5; > Chapter 39, Vol. 1, Part 6) such as alkylresorcinols and alkylpyrones, are encountered in the membrane of dormant cysts of the

Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions

27

. Figure 4 Biosynthesis of ornithine-containing lipids (for details see text).

Gram-negative bacterium Azotobacter vinelandii. Remarkably, a type I fatty acid synthase composed of ArsA and ArsD forms C22-C26 fatty acyl residues from malonyl-CoA (Miyanaga et al., 2008). These long-chain acyl residues are then used by the type III polyketide synthases ArsB or ArsC to in condensation reactions with 2–3 malonyl-CoA molecules to form alkylresorcinols or alkylpyrones, respectively.

8

Conclusions and Research Needs

Membrane lipids act in at least two ways. On one hand they form the lipid bilayer surrounding every cell and on the other hand they interact with other biomolecules based on their distinct chemical nature. Bacteria produce an unusually wide spectrum of different membrane lipid classes and for many of these lipids, important aspects on the genetics, biochemistry, and functionality are still unknown. Getting to a profound understanding of membrane protein functionality represents another bottleneck in biological research. Frequently, the function and crystallizability (in order to obtain structures) of membrane proteins depend on specific membrane lipid components and therefore makes it also an imperative from this viewpoint to study bacterial membrane lipids in much more detail. Certain essential membrane lipids are made by pathways harboring enzymes unique to bacteria. Detecting specific inhibitors for these enzymes should lead to the development of next generation antibiotics. Finally,

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membrane lipids of the bacterial cell surface, such as lipopolysaccharides, sphingolipids, ornithine-containing lipids, phosphatidylinositol-derived lipids, are of crucial importance for the interaction with eukaryotic hosts and therefore key to understand symbiotic or pathogenic processes.

References Bara´k I, Muchova´ K, Wilkinson AJ, O’Toole PJ, Pavlendova´ N (2008) Lipid spirals in Bacillus subtilis and their role in cell division. Mol Microbiol 68: 1315–1327. Benning C, Beatty JT, Prince RC, Somerville CR (1993) The sulfolipid sulfoquinovosyldiacylglycerol is not required for photosynthetic electron transport in Rhodobacter sphaeroides but enhances growth under phosphate limitation. Proc Natl Acad Sci USA 90: 1561–1565. Benning C (1998) Biosynthesis and function of the sulfolipid sulfoquinovosyldiacylglycerol. Annu Rev Plant Physiol Plant Mol Biol 49: 53–75. Benning C (2007) Questions remaining in sulfolipid biosynthesis: a historical perspective. Photosynth Res 92: 199–203. Berry AM, Harriott OT, Moreau RA, Osman SF, Benson DR, Jones AD (1993) Hopanoid lipids compose the Frankia vesicle envelope, presumptive barrier of oxygen diffusion to nitrogenase. Proc Natl Acad Sci USA 90: 6091–6094. Conover GM, Martinez-Morales F, Heidtman ML, Luo ZQ, Tang M, Chen C, Geiger O, Isberg RR (2008) Phosphatidylcholine synthesis is required for optimal function of Legionella pneumophila virulence determinants. Cell Microbiol 10: 514–528. Dowhan W, Bogdanov M, Mileykovskaya E (2008) Functional roles of lipids in membranes. In Biochemistry of lipids, lipoproteins and membranes, 5th edn. DE Vance, JE Vance (eds.). Amsterdam: Elsevier, pp. 1–37. Gao JL, Weissenmayer B, Taylor AM, Thomas-Oates J, Lo´pez-Lara IM, Geiger O (2004) Identification of a gene required for the formation of lyso-ornithine lipid, an intermediate in the biosynthesis of ornithine-containing lipids. Mol Microbiol 53: 1757–1770. Gibbons HS, Reynolds CM, Guan Z, Raetz CRH (2008) An inner membrane dioxygenase that generates the 2-hydroxymyristate moiety of Salmonella lipid A. Biochemistry 47: 2814–2825. Hacker S, Sohlenkamp C, Aktas M, Geiger O, Narberhaus F (2008) Multiple phospholipid N-methyltransferases with distinct substrate specificities are encoded in Bradyrhizobium japonicum. J Bacteriol 190: 571–580.

Hoffmann C, Leis A, Niederweis M, Plitzko JM, Engelhardt H (2008) Disclosure of the mycobacterial outer membrane. Proc Natl Acad Sci USA 105: 3963–3967. Jackson M, Crick DC, Brennan PJ (2000) Phosphatidylinositol is an essential phospholipid in mycobacteria. J Biol Chem 275: 30092–30099. Jorasch P, Wolter FP, Za¨hringer U, Heinz E (1998) A UDP glucosyltransferase from Bacillus subtilis successively transfers up to four glucose residues to 1,2diacylglycerol: expression of ypfP in Escherichia coli and structural analysis of its reaction products. Mol Microbiol 29: 419–430. Kato M, Muto Y, Tanaka-Bandoh K, Watanabe K, Ueno K (1995) Sphingolipid composition in Bacteroides species. Anaerobe 1: 135–139. Kent C, Gee P, Lee SY, Bian X, Fenno JC (2004) A CDPcholine pathway for phosphatidylcholine biosynthesis in Treponema denticola. Mol Microbiol 51: 471–481. ¨ jemyr L, Tagscherer KE, Widmalm G, Klement MLR, O Wieslander A˚ (2007) A processive lipid glycosyltransferase in the small human pathogen Mycoplasma pneumoniae: involvement in host immune response. Mol Microbiol 65: 1444–1457. Lo´pez-Lara IM, Sohlenkamp C, Geiger O (2003) Membrane lipids in plant-associated bacteria: their biosyntheses and possible functions. Mol Plant Microbe Interact 16: 567–579. Lu YJ, Zhang YM, Grimes KD, Qi J, Lee RE, Rock CO (2006) Acyl-phosphates initiate membrane phospholipid synthesis in Gram-positive pathogens. Mol Cell 23: 765–772. Matsumoto K, Kusaka J, Nishibori A, Hara H (2006) Lipid domains in bacterial membranes. Mol Microbiol 61: 1110–1117. Michell RH (2008) Inositol derivatives: evolution and functions. Nature Rev Mol Cell Biol 9: 151–161. Miyanaga A, Funa N, Awakawa T, Horinouchi S (2008) Direct transfer of starter substrates from type I fatty acid synthase to type III polyketide synthases in phenolic lipid synthesis. Proc Natl Acad Sci USA 105: 871–876. Nelson DL, Cox MM (2008) Lehninger – Principles of Biochemistry, 5th edition. New York: WH Freeman and Company.

Formation of Bacterial Membrane Lipids: Pathways, Enzymes, Reactions Poralla K, Muth G, Ha¨rtner T (2000) Hopanoids are formed during transition from substrate to aerial hyphae in Streptomyces coelicolor A3(2). FEMS Microbiol Lett 189: 93–95. Raetz CRH (1986) Molecular genetics of membrane phospholipids synthesis. Annu Rev Genet 20: 253–295. Raetz CR, Reynolds CM, Trent MS, Bishop RE (2007) Lipid A modification systems in Gram-negative bacteria. Annu Rev Biochem 76: 295–329. Riekhof WR, Andre C, Benning C (2005) Two enzymes, BtaA and BtaB, are sufficient for betaine lipid biosynthesis in bacteria. Arch Biochem Biophys 441: 96–105. Rock CO (2008) Fatty acids and phospholipids metabolism in prokaryotes. In Biochemistry of Lipids, Lipoproteins and Membranes, 5th edn. DE Vance, JE Vance (eds.). Amsterdam: Elsevier, pp. 59–96. Rojas-Jime´nez K, Sohlenkamp C, Geiger O, Martı´nezRomero E, Werner D, Vinuesa P (2005) A ClC chloride channel homologue and ornithinecontaining membrane lipids of Rhizobium tropici CIAT899 are involved in symbiotic efficiency and acid tolerance. Mol Plant Microbe Interact 18: 1175–1185. Romantsov T, Stalker L, Culham DE, Wood JM (2008) Cardiolipin controls the osmotic stress response and the subcellular location of transporter ProP in Escherichia coli. J Biol Chem 283: 12314–12323.

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Roy H, Ibba M (2008) RNA-dependent lipid remodeling by bacterial multiple peptide resistance factors. Proc Natl Acad Sci USA 105: 4667–4672. Sohlenkamp C, Galindo-Lagunas KA, Guan Z, Vinuesa P, Robinson S, Thomas Oates J, Raetz CRH, Geiger O (2007) The lipid lysyl-phosphatidylglycerol is present in membranes of Rhizobium tropici CIAT899 and confers increased resistance to polymyxin B under acidic growth conditions. Mol Plant Microbe Interact 20: 1421–1430. Sohlenkamp C, Lo´pez-Lara IM, Geiger O (2003) Biosynthesis of phosphatidylcholine in bacteria. Prog Lipid Res 42: 115–162. Summons RE, Bradley AS, Jahnke LL, Waldbauer JR (2006) Steroids, triterpenoids and molecular oxygen. Phil Trans R Soc B 361: 951–968. Wu D, Zajonc DM, Fujio M, Sullivan BA, Kinjo Y, Kronenberg M, Wilson IA, Wong CH (2006) Design of natural killer T cell activators: Structure and function of a microbial glycosphingolipid bound to mouse CD1d. Proc Natl Acad Sci USA 103: 3972–3977. Yard BA, Carter LG, Johnson KA, Overton IM, Dorward M, Liu H, McMahon SA, Oke M, Puech D, Barton GJ, Naismith JH, Campopiano DJ (2007) The structure of serine palmitoyltransferase; gateway to sphinglipid biosynthesis. J Mol Biol 370: 870–886.

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28 Lipid A R. E. Bishop Department of Biochemistry and Biomedical Sciences, McMaster University, Hamilton, ON, Canada Health Sciences Centre, Hamilton, ON, Canada [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410 2 Endotoxin Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 410 3 The Outer Membrane Permeability Barrier . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 412 4 Lipid A Modifications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 412 5 L-Ara4N Cluster . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 412 6 EptA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 414 7 PagP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 414 8 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 414

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_28, # Springer-Verlag Berlin Heidelberg, 2010

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Lipid A

Abstract: Diverse lipid A structures have been observed in a multitude of Gram-negative bacteria, but the metabolic logic of lipid A biosynthesis is widely conserved. This chapter will start by describing the nine constitutive enzymes of the Raetz pathway, which catalyze conserved lipid A biosynthetic reactions that depend on cytoplasmic cofactors. Concomitant with lipid A export and assembly on the cell surface, a number of regulated covalent modifications of lipid A can occur in the extracytoplasmic compartments. The narrow phylogenetic distribution of the lipid A modification enzymes, combined with the diverse regulatory signals governing their expression, is responsible for most of the lipid A structural diversity that is observed in nature. By focusing on E. coli as a model system, the general principles of lipid A biosynthesis and assembly are revealed to inform related processes that occur in more divergent organisms.

1

Introduction

In 1892, Richard Pfeiffer first defined endotoxin as a heat stable toxic substance that was released upon disruption of microbial envelopes (Beutler and Rietschel, 2003). The bioactive lipid A component of lipopolysaccharide (LPS) is arguably the most potent of the substances that fit Pfeiffer’s endotoxin definition. Gram-negative bacteria can modulate the structure of lipid A in order to evade detection by the host immune system (Raetz et al., 2007). This chapter summarizes the recently elucidated pathways for the biosynthesis of lipid A in Escherichia coli, which provides a framework for understanding lipid A structure and function in all Gram-negative bacteria.

2

Endotoxin Biosynthesis

The molecular genetics and enzymology of the conserved steps of lipid A biosynthesis are best characterized in E. coli, as shown in > Fig. 1. The Raetz pathway begins with the key precursor molecule UDP-GlcNAc, which is also the first substrate for peptidoglycan biosynthesis (Raetz and Whitfield, 2002). The first enzyme in lipid A biosynthesis is a cytoplasmic acyltransferase LpxA, which selectively transfers thiolester-activated 3-OH-14:0 from acyl carrier protein (ACP) to the 3-OH of UDP-GlcNAc. The crystal structure of LpxA revealed a homotrimeric molecule, which self-associates by a distinctive left handed parallel b-helix motif (Raetz and Roderick, 1995). E. coli LpxA is extraordinarily selective for 3-OH-14:0-ACP as the acyl donor substrate while the Pseudomonas aeruginosa LpxA prefers 3-OH-10:0-ACP. However, mutating certain key residues lining the active site cleft could modulate acyl chain selection. For example, the specificity for the G173M mutant of E. coli LpxA was shifted to 3-OH-10:0-ACP. In contrast, the specificity of P. aeruginosa LpxA could be extended to accommodate 3-OH-14:0-ACP in the corresponding M169G mutant (Wyckoff et al., 1998). These findings demonstrated the existence of precise hydrocarbon rulers in LpxA’s, which can explain variations in lipid A acylation that are observed between different organisms. The acylation of UDP-GlcNAc by LpxA is thermodynamically unfavorable, and the first committed step in lipid A biosynthesis is the subsequent deacetylation catalyzed by LpxC (EnvA). LpxC is a Zn2+-dependent enzyme that is an established target for antibiotic development (Onishi et al., 1996). The crystal and NMR structures of Aquifex LpxC revealed two slightly different models for the mechanism of catalysis, but both include a critical role for Zn2+

Lipid A

28

. Figure 1 The Raetz pathway for synthesis of Kdo2-lipid A. LpxA catalyzes the addition of 3-OH-14:0 to position 3 of UDP-GlcNAc. LpxC then removes the acetamido group at position 2, which allows LpxD to add a second 3-OH-14:0 group. LpxH cleaves the nucleotide to generate lipid X, which is then condensed with UDP-diacyl-GlcN to generate the disaccharide 1-phosphate. The 40 -kinase LpxK then generates lipid IVA, which is converted into Kdo2-lipid IVA by a bifunctional Kdo transferase KdtA. Kdo2-lipid IVA is a substrate for the LpxL and LpxM acyltransferases, which generate the acyloxyacyl linkages at positions 20 and 30 , respectively.

(Coggins et al., 2003; Whittington et al., 2003). Most LpxC inhibitors are hydroxamate compounds that interact with the catalytic Zn2+-ion. Current challenges are aimed at the development of inhibitors with the ability to evade efflux pumps that provide resistance, particularly in pseudomonads. Following deacetylation, an N-linked 3-OH-14:0 moiety is incorporated from ACP by LpxD (FirA) to generate UDP-2,3-diacylglucosamine (Kelly et al., 1993). A highly selective pyrophosphatase LpxH then cleaves UDP-2,3-diacylglucosamine to form lipid X (Babinski et al., 2002). Next the disaccharide synthase, LpxB, condenses UDP-2,3-diacylglucosamine and lipid X to generate the b-10 ,6-linkage found in all lipid A molecules. The membrane-bound 40 kinase LpxK then phosphorylates the disaccharide 1-phosphate to produce lipid IVA, which is an important pharmacological agent because it functions as an endotoxin antagonist in human cell lines (Garrett et al., 1997). Next, two 3-deoxy-D-manno-2-octulosonic acid (Kdo) sugars are incorporated by a Kdo transferase, which is encoded by the kdtA (waaA) gene, using the labile nucleotide CMP-Kdo as the Kdo donor. The final lipid A biosynthetic steps that

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Lipid A

occur on the cytoplasmic side of the inner membrane depend on the prior addition of the Kdo sugars and involve the transfer of lauroyl (12:0) and myristoyl (14:0) groups from ACP to the distal glucosamine unit to produce acyloxyacyl linkages; the reactions are catalyzed at the 20 position by LpxL (HtrB) and at the 30 -position by LpxM (MsbB), respectively. Under conditions of cold growth at 12 C, LpxL is replaced by LpxP, which has a preference for palmitoleate (16:1cisD9) in ACP (Carty et al., 1999). The incorporation of an unsaturated acyl chain into lipid A likely increases membrane fluidity under cold growth conditions. Viable mutants that lack acyloxyacyl linkages in lipid A are attenuated for virulence and reveal the importance of the lipid A acylation pattern in inflammation (Vorachek-Warren et al., 2002). All other enzymatic steps of the Raetz pathway, and those for the biosynthesis of CMP-Kdo, are essential for cell viability and, thus, provide potential targets for antibiotic development.

3

The Outer Membrane Permeability Barrier

LPS contains phosphate and acidic sugars and is therefore negatively charged. In order to reduce the electrostatic repulsion between LPS molecules at the cell surface, the bacterial outer membrane (OM) sequesters divalent cations, mainly Mg+2, which neutralize the negative charges and maintain the integrity of the OM. The presence of hydrogen-bond donors and acceptors in the lipid A molecule allows for additional lateral interactions that cannot occur between phospholipid molecules (Nikaido, 2003). Moreover, the six or seven saturated acylchains of lipid A serve to reduce the fluidity of the OM bilayer compared with the inner membrane. The tight lateral interactions between LPS, combined with low membrane fluidity, provide a permeability barrier in the OM to lipophilic solutes and detergents. However, cationic antimicrobial peptides (CAMPs) can displace Mg+2-ions from the cell surface to promote their uptake by the bacterial cell (Hancock et al., 1995).

4

Lipid A Modifications

Considering the importance of Mg+2 in maintaining the OM permeability barrier, it is not surprising that Mg+2-limitation and CAMPs can regulate the covalent structure of lipid A by triggering the PhoP/PhoQ and PmrA/PmrB signal transduction pathways (Bader et al., 2005; Groisman, 2001). Several covalent modifications of lipid A have been characterized in E. coli (> Fig. 2). PagP is a transacylase that incorporates a palmitate chain at position 2 (Bishop et al., 2000). Moreover, the phosphate groups at positions 1 and 40 of the lipid A disaccharide backbone can be modified with 4-amino-4-deoxy-L-arabinose (L-Ara4N) and/or phosphoethanolamine (pEtN), which serve to reduce the overall negative charge of lipid A (Gunn et al., 1998). Lipid A modifications occur in the extracytoplasmic compartments and help to restore the OM permeability barrier and provide resistance to CAMPs.

5

L-Ara4N Cluster

PmrA/PmrB is only one of several clusters of pmr genes that were originally identified in polymyxin resistant mutants of E. coli. The pmrF (pbgP) locus encodes an operon of 7 open reading frames pmrHFIJKLM, which, together with the unlinked pmrE (ugd), are required for

Lipid A

28

. Figure 2 Regulated covalent lipid A modifications in E. coli. The conserved lipid A nucleus can be modified by the addition of L-Ara4N and pEtN to the phosphate groups, and by the addition of a palmitate chain at position 2. Palmitoylation of lipid A is under the direct control of PhoP/PhoQ, while PmrA/PmrB controls the phosphate modifications.

L-Ara4N synthesis (Gunn et al., 1998). The first step involves the conversion of UDP-glucose into UDP-glucuronic acid catalyzed by a dehydrogenase encoded by pmrE. Complex regulation of dehydrogenase gene expression reflects the fact that UDP-glucuronic acid is a precursor for both colanic acid-containing capsules and L-Ara4N. Next, PmrI (ArnA) catalyzes the oxidative decarboxylation of UDP-glucuronic acid to generate a novel UDP-4-keto-pyranose intermediate (Breazeale et al., 2002). PmrH (ArnB) then catalyzes a transamination reaction using glutamate as the amine donor to generate UDP-L-Ara4N (Breazeale et al., 2003). The crystal structure of PmrH has verified that a pyridoxal phosphate cofactor contributes to the catalytic mechanism (Noland et al., 2002). Interestingly, PmrI contains a second domain that formylates the 4-amine of UDP-L-Ara4N (Breazeale et al., 2005). The resultant UDP-L-Ara4-Formyl-N is transferred by PmrF (ArnC) to the membrane-anchored undecaprenyl phosphate, forming undecaprenyl phosphate-L-Ara4-Formyl-N. The formylation step may drive forward the equilibrium of the transamination step, which is thermodynamically unfavorable. The necessity of a subsequent deformylation step catalyzed by PmrJ (ArnD) is dictated by the fact that undecaprenyl phosphate-L-Ara4N is the substrate for PmrK (ArnT), which catalyzes the final transfer of L-Ara4N to lipid A at the periplasmic surface of the inner membrane (Trent et al., 2001a, b). Roles for PmrL and PmrM (ArnE and ArnF) in flipping

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undecaprenyl phosphate-L-Ara4N across the inner membrane have been recently confirmed (Yan et al., 2007).

6

EptA

The pEtN adding enzyme EptA has been cloned from E. coli (Lee et al., 2004), and a homologous gene from Neisseria has been associated with the addition of pEtN to lipid A (Cox et al., 2003). The EptA-encoding gene is the upstream open reading frame that is part of the pmrAB operon, and is also known as pmrC (pagB). The EptA active site is located on the periplasmic side of the inner membrane. Phosphatidylethanolamine is the reported pEtN donor and several EptA homologues are likely responsible for pEtN addition to other cell envelope components including the inner core sugars of LPS (Reynolds et al., 2005). It is noteworthy that roughly one third of E. coli lipid A carries a diphosphate moiety instead of the monophosphate at position 1 (Touze´ et al., 2008), and that the responsible phosphorylating enzyme shares with EptA the ability to generate a phosphodiester bond at the same position in lipid A.

7

PagP

PagP functions to transfer a palmitate chain from a phospholipid to the hydroxyl group of the N-linked 3-OH-14:0 chain on the proximal glucosamine unit of lipid A (Bishop, 2005). PagP is regulated by PhoP/PhoQ and was the first enzyme of lipid A biosynthesis shown to be localized in the OM (Bishop et al., 2000; Jia et al., 2004). Since thiolester-containing substrates are not available in the extracellular compartments, PagP uses a phospholipid as the palmitoyl donor instead. PagP was first identified in the salmonellae due to its role in providing resistance to cationic antimicrobial peptides (Guo et al., 1998), and was subsequently purified from E. coli (Bishop et al., 2000). In addition to these enteric pathogens, PagP homologues are present in the respiratory pathogens Legionella pneumophila and Bordetella bronchiseptica, where PagP has been shown to be necessary for disease-causation in animal models of infection (Preston et al., 2003; Robey et al., 2001). PagP homologues are also found in Yersinia, Photorhabdus and Erwinia species, which adopt pathogenic lifestyles in animals, insects, and plants, respectively. The solution and crystal structures of PagP indicate that the enzyme is activated in the OM in response to perturbations of lipid asymmetry (Ahn et al., 2004; Hwang et al., 2002). Phospholipids must gain access to the PagP hydrocarbon ruler after first migrating into the OM external leaflet (Khan et al., 2007). Recent evidence indicates PagP can function as a sensory transducer, which is triggered by perturbations to OM lipid asymmetry (Smith et al., 2008).

8

Research Needs

Lipid A and its regulated covalent modifications exhibit profound effects on bacterial and human physiology. Novel endotoxin antagonists and immune adjuvants have already been developed from modified lipid A structures (Christ et al., 1995; Ulrich and Myers, 1995). By revealing the biochemical details of lipid A structure and function we hope to understand its role in bacterial pathogenesis and to intervene with novel treatments for infection.

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However, we must remind ourselves that multiple molecular subtypes of lipid A are acting in concert in the bacterial cell. The need to unravel the interactions between individual lipid A modifications will provide fertile ground for future research.

Acknowledgments Work in the author’s laboratory was supported by the Canadian Institutes of Health Research. We apologize to those authors whose work could not be cited due to space limitations.

References Ahn VE, Lo EI, Engel CK, Chen L, Hwang PM, Kay LE, Bishop RE, Prive´ GG (2004) A hydrocarbon ruler measures palmitate in the enzymatic acylation of endotoxin. EMBO J 23: 2931–2941. Babinski KJ, Kanjilal SJ, Raetz CR (2002) Accumulation of the lipid A precursor UDP-2,3-diacylglucosamine in an Escherichia coli mutant lacking the lpxH gene. J Biol Chem 277: 25947–25956. Bader MW, Sanowar S, Daley ME, Schneider AR, Cho U, Xu W, Klevit RE, Le Moual H, Miller SI (2005) Recognition of antimicrobial peptides by a bacterial sensor kinase. Cell 122: 461–472. Beutler B, Rietschel ET (2003) Innate immune sensing and its roots: the story of endotoxin. Nat Rev Immunol 3: 169–117. Bishop RE (2005) The lipid A palmitoyltransferase PagP: molecular mechanisms and role in bacterial pathogenesis. Mol Microbiol 57: 900–912. Bishop RE, Gibbons HS, Guina T, Trent MS, Miller SI, Raetz CR (2000) Transfer of palmitate from phospholipids to lipid A in outer membranes of Gramnegative bacteria. EMBO J 19: 5071–5080. Breazeale SD, Ribeiro AA, McClerren AL, Raetz CR (2005) A formyltransferase required for polymyxin resistance in Escherichia coli and the modification of lipid A with 4-amino-4-deoxy-L-arabinose: identification and function of UDP-4-deoxy-4-formamidoL-arabinose. J Biol Chem 280: 14154–14167. Breazeale SD, Ribeiro AA, Raetz CR (2002) Oxidative decarboxylation of UDP-glucuronic acid in extracts of polymyxin-resistant Escherichia coli. Origin of lipid A species modified with 4-amino-4-deoxy-Larabinose. J Biol Chem 277: 2886–2896. Breazeale SD, Ribeiro AA, Raetz CR (2003) Origin of lipid A species modified with 4-amino-4-deoxyL-arabinose in polymyxin resistant mutants of Escherichia coli: an aminotransferase (ArnB) that generates UDP-4-amino-4-deoxy-L-arabinose. J Biol Chem 278: 24731–24739.

Carty SM, Sreekumar KR, Raetz CR (1999) Effect of cold shock on lipid A biosynthesis in Escherichia coli. Induction at 12 degrees C of an acyltransferase specific for palmitoleoyl-acyl carrier protein. J Biol Chem 274: 9677–9685. Christ WJ, Asano O, Robidoux AL, Perez M, Wang YA, Dubuc GR, Gavin WE, Hawkins LD, McGuinness PD, Mullarkey MA, Lewis MD, Kishi Y, Kawata T, Bristol JR, Rose JR, Rossignol DP, Kobayashi S, Hishinuma L, Kimura A, Asakawa N, Katayama K, Yamatsu I (1995) E5531, a pure endotoxin antagonist of high potency. Science 268: 80–83. Coggins BE, Li X, McClerren AL, Hindsgaul O, Raetz CR, Zhou P (2003) Structure of the LpxC deacetylase with a bound substrate-analog inhibitor. Nat Struct Biol 10: 645–651. Cox AD, Wright JC, Li J, Hood DW, Moxon ER, Richards JC (2003) Phosphorylation of the lipid a region of meningococcal lipopolysaccharide: identification of a family of transferases that add phosphoethanolamine to lipopolysaccharide. J Bacteriol 185: 3270–3277. Garrett TA, Kadrmas JL, Raetz CR (1997) Identification of the gene encoding the Escherichia coli lipid A 40 -kinase. Facile phosphorylation of endotoxin analogs with recombinant LpxK. J Biol Chem 272: 21855–21864. Groisman EA (2001) The pleiotropic two-component regulatory system PhoP-PhoQ. J Bacteriol 183: 1835–1842. Gunn JS, Lim KB, Krueger J, Kim K, Guo L, Hackett M, Miller SI (1998) PmrA-PmrB-regulated genes necessary for 4-aminoarabinose lipid A modification and polymyxin resistance. Mol Microbiol 27: 1171–1182. Guo L, Lim K, Poduje C, Daniel M, Gunn J, Hackett J, Miller SI (1998) Lipid A acylation and bacterial resistance against vertebrate anti-microbial peptides. Cell 95: 189–198.

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Hancock RE, Falla T, Brown M (1995) Cationic bactericidal peptides. Adv Microb Physiol 37: 135–175. Hwang PM, Choy WY, Lo EI, Chen L, Forman-Kay JD, Raetz CR, Prive´ GG, Bishop RE, Kay LE (2002) Solution structure and dynamics of the outer membrane enzyme PagP by NMR. Proc Natl Acad Sci USA 99: 13560–13565. Jia W, El Zoeiby A, Petruzziello TN, Jayabalasingham B, Seyedirashti S, Bishop RE (2004) Lipid trafficking controls endotoxin acylation in outer membranes of Escherichia coli. J Biol Chem 279: 44966–44975. Kelly TM, Stachula SA, Raetz CR, Anderson MS (1993) The firA gene of Escherichia coli encodes UDP-3-O-(R-3-hydroxymyristoyl)-glucosamine Nacyltransferase. The third step of endotoxin biosynthesis. J Biol Chem 268: 19866–19874. Khan MA, Neale C, Michaux C, Pome`s R, Prive´ GG, Woody RW, Bishop RE (2007) Gauging a hydrocarbon ruler by an intrinsic exciton probe. Biochemistry 46: 4565–4579. Lee H, Hsu FF, Turk J, Groisman EA (2004) The PmrAregulated pmrC gene mediates phosphoethanolamine modification of lipid A and polymyxin resistance in Salmonella enterica. J Bacteriol 186: 4124–4133. Nikaido H (2003) Molecular basis of bacterial outer membrane permeability revisited. Microbiol Mol Biol Rev 67: 593–656. Noland BW, Newman JM, Hendle J, Badger J, Christopher JA, Tresser J, Buchanan MD, Wright TA, Rutter ME, Sanderson WE, Muller-Dieckmann HJ, Gajiwala KS, Buchanan SG (2002) Structural studies of Salmonella typhimurium ArnB (PmrH) aminotransferase: a 4-amino-4-deoxy-L-arabinose lipopolysaccharide-modifying enzyme. Structure 10: 1569–1580. Onishi HR, Pelak BA, Gerckens LS, Silver LL, Kahan FM, Chen MH, Patchett AA, Galloway SM, Hyland SA, Anderson MS, Raetz CR (1996) Antibacterial agents that inhibit lipid A biosynthesis. Science 274: 980–982. Preston A, Maxim E, Toland E, Pishko EJ, Harvill ET, Caroff M, Maskell DJ (2003) Bordetella bronchiseptica PagP is a Bvg-regulated lipid A palmitoyl transferase that is required for persistent colonization of the mouse respiratory tract. Mol Microbiol 48: 725–736. Raetz CR, Reynolds CM, Trent MS, Bishop RE (2007) Lipid A modification systems in gram-negative bacteria. Annu Rev Biochem 76: 295–329. Raetz CR, Roderick SL (1995) A left-handed parallel beta helix in the structure of UDP-N-acetylglucosamine acyltransferase. Science 270: 997–1000. Raetz CR, Whitfield C (2002) Lipopolysaccharide endotoxins. Annu Rev Biochem 71: 635–700.

Reynolds CM, Kalb SR, Cotter RJ, Raetz CR (2005) A phosphoethanolamine transferase specific for the outer 3-deoxy-D-manno-octulosonic acid residue of Escherichia coli lipopolysaccharide. Identification of the eptB gene and Ca2+ hypersensitivity of an eptB deletion mutant. J Biol Chem 280: 21202–21211. Robey M, O’Connell W, Cianciotto NP (2001) Identification of Legionella pneumophila rcp, a pagP-like gene that confers resistance to cationic antimicrobial peptides and promotes intracellular infection. Infect Immun 69: 4276–4286. Smith AE, Kim SH, Liu F, Jia W, Vinogradov E, Gyles CL, Bishop RE (2008) PagP activation in the outer membrane triggers R3 core oligosaccharide truncation in the cytoplasm of Escherichia coli O157:H7. J Biol Chem 283: 4332–4343. Trent MS, Ribeiro AA, Doerrler WT, Lin S, Cotter RJ, Raetz CR (2001a) Accumulation of a polyisoprenelinked amino sugar in polymyxin-resistant Salmonella typhimurium and Escherichia coli: structural characterization and transfer to lipid A in the periplasm. J Biol Chem 276: 43132–43144. Trent MS, Ribeiro AA, Lin S, Cotter RJ, Raetz CR (2001b) An inner membrane enzyme in Salmonella and Escherichia coli that transfers 4-amino-4-deoxy-Larabinose to lipid A: induction on polymyxinresistant mutants and role of a novel lipid-linked donor. J Biol Chem 276: 43122–43131. Touze´ T, Tran AX, Hankins JV, Mengin-Lecreulx D, Trent MS (2008) Periplasmic phosphorylation of lipid A is linked to the synthesis of undecaprenyl phosphate. Mol Microbiol 67: 264–277. Ulrich JT, Myers KR (1995) Monophosphoryl lipid A as an adjuvant. In Vaccine Design: The Subunit and Adjuvant Approach. MF Powell and MJ Newman (eds.). New York: Plenum Press, pp. 495–524. Vorachek-Warren MK, Ramirez S, Cotter RJ, Raetz CR (2002) A triple mutant of Escherichia coli lacking secondary acyl chains on lipid A. J Biol Chem 277: 14194–14205. Whittington DA, Rusche KM, Shin H, Fierke CA, Christianson DW (2003) Crystal structure of LpxC, a zinc-dependent deacetylase essential for endotoxin biosynthesis. Proc Natl Acad Sci USA 100: 8146–8150. Wyckoff TJ, Lin S, Cotter RJ, Dotson GD, Raetz CR (1998) Hydrocarbon rulers in UDP-N-acetylglucosamine acyltransferases. J Biol Chem 273: 32369–32372. Yan A, Guan Z, Raetz CR (2007) An undecaprenyl phosphate-aminoarabinose flippase required for polymyxin resistance in Escherichia coli. J Biol Chem 282: 36077–36089.

29 Membrane Biogenesis H. Goldfine Department of Microbiology, University of Pennsylvania School of Medicine, Philadelphia, PA, USA [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418

2 2.1 2.2 2.3

Lipid Bilayer Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418 Requirements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 418 The Liquid Crystalline Phase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 419 The Formation of Non-Bilayer Phases: Effects of Hydrocarbons, Solvents, and Other Small Molecules . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 420

3

Membrane Vesicle Shedding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422

4

Lipid Domains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 422

5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 423

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_29, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: In order to maintain a fluid lipid bilayer in the cell membrane, microorganisms must adjust to environmental conditions including the ambient temperature, pressure, and the presence of solutes that affect the physical state of the membrane. Although the types of amphipathic lipids present in the cell membrane can vary widely between species, the types of adjustments made, including changes in the compositions of the hydrocarbon chains and the polar head groups, appear to obey certain rules, which are discussed in this chapter. Abbreviations: DGDG, diglycosyldiacylglycerol; GAPlaE, glycerol acetal of the plasmalogen form of phosphatidylethanolamine; MGDG, monoglycosyldiacylglycerol; MV, membrane vesicles; PE, phosphatidylethanolamine; PlaE, plasmalogen form of phosphatidylethanolamine; PG, phosphatidylglycerol

1

Introduction

Biological membranes are self-assembled structures largely composed of lipids and proteins. The cell membrane forms a semipermeable barrier and contains proteins involved in solute transport, the formation of chemical energy, the synthesis and translocation of macromolecules, and the synthesis of the membrane lipids. Microbes are no exception, and they show considerable diversity with respect to the types of lipids they contain. Of paramount importance for the formation of a stable, functional membrane is the ability of the ensemble of lipids to form a fluid bilayer within the growth temperature boundaries of the organism. In most cases the predominant polar lipids are phospholipids, but in some organisms glycosyldiacylglycerols represent a significant fraction of the total membrane lipids. In this chapter the general rules for fluid bilayer formation and maintenance will be considered as will the special requirements of organisms that produce apolar compounds.

2

Lipid Bilayer Formation

2.1

Requirements

Most bacterial membrane lipids are amphipathic or bipolar. They have relatively long hydrocarbon chains, predominantly from 14 to 20 carbons long, which are usually, but not always, linked to a glycerol backbone by means of an ester, ether, or alk-1-enyl ether bond. The hydrocarbon chains are hydrophobic, point towards the center of the bilayer, and form stable associations with adjacent chains through hydrophobic, non-covalent interactions. At the sn-3 position of the glycerol backbone in bacteria or the sn-1 position in Archaea a polar, hydrophilic group is attached and in the case of phospholipids this group will consist of a phosphodiester to a short chain primary alcohol such as ethanolamine, glycerol, or serine and in a subgroup of bacteria choline. In the bacteria that have glycolipids these are of the glycosyl diradyl (diradyl refers to chains that may be linked through acyl ester bonds, alk-1-enyl ether or saturated ether bonds) glycerol type (> Fig. 1). In order to form a bilayer, a significant proportion of the lipid molecules must be cylindrical in shape (Israelachvili et al., 1980). These lipids will spontaneously form bilayers with minimum curvature on the scale of a cell membrane. Among the polar lipids in bacteria that tend to form bilayers are phosphatidylglycerol (PG), phosphatidylserine, and

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. Figure 1 Structures of some bacterial lipids. (a) Phosphatidylethanolamine; (b) plasmalogen form of phosphatidylethanolamine; (c) glycerol acetal of the plasmalogen form of phosphatidylethanolamine; (d) glucosyldiacylglycerol an example of a monoglycosyldiacylglycerol.

diglycosyldiacylglycerol (DGDG). In most bacterial membranes there is a significant proportion of polar lipids that do not tend to form bilayers, but rather produce the inverted hexagonal (HII) or other nonlamellar phases (> Fig. 2). These lipids are conical in shape with relatively small polar head groups and acyl chains that sweep out a wider conical volume than that occupied by the head group. Among these are phosphatidylethanolamine (PE), monoglycosyldiacylglycerols (MGDG) and cardiolipin in the presence of certain divalent metal ions (Cullis and De Kruijff, 1979). These non-bilayer-preferring lipids are normally present with sufficient bilayer-forming lipids to maintain the bilayer arrangement.

2.2

The Liquid Crystalline Phase

At their normal growth temperatures, biological membranes are said to be fluid or to be more precise, to exist in the lamellar liquid crystalline phase, termed La. In this state the hydrocarbon chains are disordered with rapid flexion all along the chains. The intact lipid molecules rotate in the plane of the bilayer and diffuse laterally. They rarely flip from one side of the bilayer to the other unless aided by proteins called flipases or phospholipid translocators. This state is essential for the normal function of membrane proteins embedded within the bilayer. If the ambient temperature cools, lipids in the bilayer will eventually undergo a transition to a gel (Lb) phase in which the chains are packed in a stiff, near crystalline arrangement. Nature ensures the fluidity of bacterial membranes by incorporating a significant proportion of either unsaturated or branched hydrocarbon chains in the lipids. In general the former is true

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. Figure 2 A typical bilayer and an illustration of the tube-like structure of the hexagonal II phase in which the center is aqueous. These tubes are packed into hexagonal arrays in which the tips of the hydrocarbon chains are touching.

for gram negative bacteria while the latter is the case in gram positive bacteria, but these rules do not hold universally. Unsaturation in hydrocarbon chains introduces a kink which decreases lateral close packing, thus lowering the temperature of transition to a gel state. The presence of methyl branches usually at the penultimate (anteiso) or the terminal carbon (iso) also deters close packing with a similar result. When bacteria are grown at lower temperatures they usually increase either the content of unsaturated chains or the ratio of anteiso to iso chains.

2.3

The Formation of Non-Bilayer Phases: Effects of Hydrocarbons, Solvents, and Other Small Molecules

As the ambient temperature is increased above the melting point, bilayers consisting of mixtures of polar lipids may eventually transition to other liquid crystalline states including the cubic phase Qa, and the inverted hexagonal HII phase. The temperature at which this occurs depends on the lipid polar head groups and the compositions of the hydrocarbon chains. As noted above, smaller head groups promote the formation of nonlamellar phases. Hydrogen bonding of the polar head groups can also cause them to come close together, favoring the formation of a nonlamellar phase. In addition, for example, in the case of PE or MGDG, the presence of unsaturated hydrocarbon chains increases the tendency to form nonlamellar phases (Cullis and De Kruijff, 1979; Lindblom and Rilfors, 1989). The lateral pressure exerted by these lipids in the bilayer leads to curvature stress. Near the lamellar– nonlamellar transition, ‘‘The bilayer leaflets are literally straining to bend’’ (Gruner and Shyamsunder, 1991). Bacteria have adopted different strategies to avoid the disruption of the bilayer caused by formation of non-bilayer arrangements. For example, when the degree of unsaturation of the membrane lipids in Acholeplasma laidlawii is increased artificially the ratio of diglucosyldiacylglycerol (DGlcDAG) to monoglucosyldiacylglycerol (MGlcDAG) increases. The ratios of other polar lipids are also changed, and the overall result is to decrease the tendency to form nonlamellar structures (Rilfors and Lindblom, 2002; Wieslander et al., 1980). In Clostridium butyricum a similar degree of enrichment with unsaturated hydrocarbons chains leads to an increase in the amount of a glycerol acetal of the plasmalogen form of PE (GAPlaE) and a decrease in diradyl PE (> Fig. 1), again leading to stabilization of

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the bilayer (Goldfine et al., 1987). Studies with PE-deficient mutants of E. coli lead to a similar conclusion: the bacteria maintain an optimal ratio between lipids that prefer the bilayer structure and those that prefer the HII phase (Rietveld et al., 1993). Furthermore, it has been shown with both A. laidlawii and E. coli that these cells appear to adjust the composition of their membrane lipids in order to maintain lamellar liquid crystalline phases, avoiding the formation of a gel phase and nonlamellar liquid crystalline phases. The cells are said to grow in a ‘‘window’’ between the boundaries of the gel and nonlamellar phases (Rilfors and Lindblom, 2002). C. butyricum appears to regulate its lipid composition according to a similar plan (Goldfine et al., 1987). Production of solvents or growth in the presence of solvents presents special problems for microorganisms that have these life styles. In A. laidlawii (Wieslander et al., 1986) and C. butyricum (MacDonald and Goldfine, 1991) growth in the presence of solvents such as cyclohexane increases the proportions of the bilayer-preferring membrane lipids DGlcDAG and GAPlaE, respectively. Shorter chain alcohols such as butanol have little effect. Shorter chain alcohols are expected to expand the headgroup region thus stabilizing the lamellar phase (Janes, 1996; Weber and De Bont, 1996). Longer chain alcohols will expand the hydrocarbon region in addition to the headgroup region and thus will stabilize the HII over the lamellar state. This is counterbalanced by increasing the proportions of bilayer-preferring lipids in both organisms. In experimental systems in which alcohols are added to PE, the crossover point is between butanol and longer chain alcohols (Weber and De Bont, 1996) and similar crossover points were seen with A. laidlawii and C. butyricum (see references above). Other adaptive changes are seen in microorganisms in response to the presence of alcohols and other solvents. These compounds increase the permeability of artificial lipid vesicles. Addition of ethanol to the growth medium of E. coli leads to an increase in the ratio of unsaturated to saturated acyl chains in the membrane lipids, whereas C6 to C10 alcohols have the opposite effect. Pseudomonas putida grown in the presence of toluene undergoes an interesting adaptation in which the normal cis-unsaturated fatty acids are converted to the corresponding trans-fatty acids. This would have the effect of increasing lipid ordering and raising the temperature of the lamellar to HII transition. Another effect seen was a decrease in the proportion of PE relative to the sum of PG and cardiolipin (See > Chapter 60, Vol. 2, Part 10), again resulting in an increase in the lamellar to HII transition temperature (Weber and De Bont, 1996). There are several solvent-producing clostridia. Among these is C. beijerinckii which has a lipid composition similar to that of C. butyricum with the exception of having phosphatidyl N-methylethanolamine in addition to PE. It also has plasmalogens and glycerol acetals of the plasmalogen forms of these lipids (Goldfine and Johnston, 2005) (> Fig. 1). Its response to artificially increased unsaturated fatty acids in the membrane lipids is similar to that described above for C. butyricum, i.e., an increase in the relative amounts of the glycerol acetals of the plasmalogens (Goldfine, 1984). The lipids of C. beijerinckii ATCC 39057, previously identified as C. acetobutylicum, were studied during acetone/butanol production. As solvents accumulated in the culture fluid, the ratio of GAPlaE + GAPlaME/diradyl PE + PME increased and the saturated/unsaturated (SU) acyl chain ratio also increased. Similar changes were seen in the acyl chains along with a shortening of these chains upon addition of n-hexanol or n-octanol. Ethanol or butanol produced smaller changes in the composition of the acyl chains (Lepage et al., 1987). These changes in polar head groups and acyl chains would tend to counteract the effects of longer chain alcohols that would perturb the lamellar/nonlamellar equilibrium as described

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above. C. acetobutylicum has a high proportion of glycosyl diradylglycerols in addition to glycerophospholipids. It responds to increased membrane lipid unsaturated chains similarly to C. beijerinckii and C. butyricum by an increase in GAPlaE relative to diradyl PE. The glycolipid composition undergoes changes similar to those seen in A. laidlawii, i.e., an increase in DGDG relative to MGDG (Johnston and Goldfine, 1992). Glycerol acetals of plasmalogens have been found in all of the solvent-producing clostridia examined to-date, but not in other clostridia (Goldfine and Johnston, 2005). All the solvent-producing clostridia have PlaE, which is more prone to undergo a lamellar to HII transition than PE. The studies cited above suggest that glycerol acetals evolved to improve the ability of these organisms to withstand the stress of solvent formation. Studies on the effects of butanol challenge and growth temperature on the membrane lipids and fluidity of C. acetobutylicum ATCC 824 and a butanol-tolerant mutant, SA-2 showed that addition of butanol to the growth medium also led to an increase in the ratio of saturated to unsaturated (plus cyclopropane) acyl chains in the membrane lipids (Baer et al., 1987; Vollherbst-Schneck et al., 1984). Shorter chain alcohols in model membranes cause a shift to lower temperatures in the transition from the gel to the liquid crystalline phase. This is often referred to as an increase in membrane fluidity, which can be measured with various probes (Weber and De Bont, 1996). The increase in the SU ratio seen in both C. beijerinckii and C. acetobutylicum upon butanol challenge would tend to counteract the effects of butanol on membrane fluidity. In summary, changes observed can be viewed as a response to the addition of solvents to the growth medium or to the production of solvents by bacteria that serves to maintain the cell membrane in an essential window (La) between the gel and nonlamellar phases.

3

Membrane Vesicle Shedding

Gram-negative bacteria are capable of shedding outer membrane vesicles (MV) by a process of blebbing. MV retain the asymmetry of the outer membrane with lipopolysaccharide facing out and glycerophospholipids facing in. Encapsulated with MV are proteins derived from the periplasmic space between the cell membrane and the outer membrane. Some of the encapsulated proteins have been found to be proteases and autolysins and the latter can serve to lyse other bacterial species in the environment (Beveridge, 1999; Mashburn-Warren and Whiteley, 2006). Since MV production has been found to be upregulated as part of the stress response, it has been proposed that the lysis function may serve to produce nutrients under limiting conditions (Mashburn-Warren and Whiteley, 2006; McBroom and Kuehn, 2007). Whether these vesicles play a role in hydrocarbon production or utilization is not known.

4

Lipid Domains

Lipid rafts or lipid microdomains are thought to exist in eukaryotic cells where they result from the segregation of cholesterol and sphingomyelin into discrete patches of membrane. Bacteria do not have cholesterol and most do not have sphingomyelin, though sterol-like, pentacyclic triterpenoids based on the hopane structure are found in many bacteria. Evidence for cardiolipin domains at the septal regions and at the poles in both Bacillus subtilis (See > Chapter 10, Vol. 1, Part 2) and E. coli based on specific staining with the fluorescent

Membrane Biogenesis

29

dye 10-N-nonyl-acridine orange has been presented (Kawai et al., 2004; Mileykovskaya and Dowhan, 2000). PE has also been found to segregate at the septal membranes of B. subtilis, but not in E. coli, in which PE is the major lipid (Nishibori et al., 2005). Based on green fluorescent protein fusions to the enzymes needed for phospholipid synthesis in B. subtilis, these authors have suggested that lipid synthesis is mostly localized in the septal membranes.

5

Research Needs

Although core knowledge on the membranes of hydrocarbon-producing bacteria is evolving, much remains to be learned about the mechanisms by which the cells respond to hydrocarbons. Is the regulation of membrane composition seen in solvent-producing bacteria the result of regulation of the production of the enzymes involved in lipid synthesis or does it result from the effects of solvents on the activities of these enzymes? What changes in the expression of membrane proteins occur (See > Chapter 45, Vol. 1, Part 7) in response to hydrocarbon production or hydrocarbons in the environment? What is the mechanism for translocation of hydrocarbons in the bacterial cell?

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Johnston NC, Goldfine H (1992) Replacement of the aliphatic chains of Clostridium acetobutylicum by exogenous fatty acids: regulation of phospholipid and glycolipid composition. J Bacteriol 174: 1848–1853. Kawai F, Shoda M, Harashima R, Sadaie Y, Hara H, Matsumoto K (2004) Cardiolipin domains in Bacillus subtilis marburg membranes. J Bacteriol 186: 1475–1483. Lepage C, Fayolle F, Hermann M, Vandecasteele J-P (1987) Changes in membrane lipid composition of Clostridium acetobutylicum during acetone-butanol fermentation: effects of solvents, growth temperature and pH. J Gen Microbiol 133: 103–110. Lindblom G, Rilfors L (1989) Cubic phases and isotropic structures formed by membrane lipids – possible biological relevance. Biochim Biophys Acta 988: 221–256. MacDonald DL, Goldfine H (1991) Effects of solvents and alcohols on the polar lipid composition of Clostridium butyricum under conditions of controlled lipid chain composition. Appl Environ Microbiol 57: 3517–3521. Mashburn-Warren LM, Whiteley M (2006) Special delivery: vesicle trafficking in prokaryotes. Mol Microbiol 61: 839–846. McBroom AJ, Kuehn MJ (2007) Release of outer membrane vesicles by gram-negative bacteria is a novel

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envelope stress response. Mol Microbiol 63: 545–558. Mileykovskaya E, Dowhan W (2000) Visualization of phospholipid domains in Escherichia coli by using the cardiolipin-specific fluorescent dye 10-N-nonyl acridine orange. J Bacteriol 182: 1172–1175. Nishibori A, Kusaka J, Hara H, Umeda M, Matsumoto K (2005) Phosphatidylethanolamine domains and localization of phospholipid synthases in Bacillus subtilis membranes. J Bacteriol 187: 2163–2174. Rietveld AG, Killian JA, Dowhan W, De Kruijff B (1993) Polymorphic regulation of membrane phospholipid composition in Escherichia coli. J Biol Chem 268: 12427–12433. Rilfors L, Lindblom G (2002) Regulation of lipid composition in biological membranes – biophysical studies of lipids and lipid synthesizing enzymes. Colloids Surf B Biointerfaces 26: 112–124.

Vollherbst-Schneck K, Sands JA, Montenecourt BS (1984) Effect of butanol on lipid composition and fluidity of Clostridium acetobutylicum ATCC 824. Appl Environ Microbiol 47: 193–194. Weber FJ, De Bont JAM (1996) Adaptation mechanisms of microorganisms to the toxic effects of organic solvents on membranes. Biochim Biophys Acta Rev Biomembr 1286: 225–245. Wieslander A˚, Christiansson A, Rilfors L, Lindblom G (1980) Lipid bilayer stability in membranes. Regulation of lipid composition in Acholeplasma laidlawii as governed by molecular shape. Biochemistry 19: 3650–3655. Wieslander A˚, Rilfors L, Lindblom G (1986) Metabolic changes of membrane lipid composition in Acholeplasma laidlawii by hydrocarbons, alcohols, and detergents: arguments for effects on lipid packing. Biochemistry 25: 7511–7517.

30 Membrane Disrupting Proteins J. H. Lakey1 . G. Anderluh2 1 Institute for Cell and Molecular Biosciences, University of Newcastle upon Tyne, Newcastle upon Tyne, UK [email protected] 2 Department of Biology, University of Ljubljana, Ljubljana, Slovenia [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 426 2 The Importance of Secondary Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 3 The Alpha-Helix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 427 4 Beta Structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 429 5 Tryptophan . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 431 6 Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 432

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Abstract: Membrane damaging proteins, found widely in biology, are extensively used by microorganisms to cause disease in hosts or attack competing organisms. The proteins must travel to their target in a water soluble state and then insert into the membrane phase. This chapter deals with the fundamental features of such proteins, highlighting the importance of secondary structure formation during membrane insertion and the role of tryptophan residues in interfacial binding.

1

Introduction

Proteins that damage membranes are generally involved in either the promotion of, or inhibition of, pathogenesis in its broadest sense. From bacteria to bees and sea anemones to mammals, membrane damage is used for attack (See > Chapter 68, Vol. 4, Part 7; > Chapter 69, Vol. 3, Part 7; > Chapter 70, Vol. 3, Part 7) and defense (Anderluh and Lakey, 2008). Rearrangement of eukaryotic membranes during normal cell turnover or their destruction during programmed cell death may also use similar mechanisms (See > Chapter 21, Vol. 2, Part 4) but these are less well understood and also not of direct relevance to this review. Whatever the reason for their evolution, the proteins’ main target is to disrupt the impermeability of the cell membrane (Anderluh and Lakey, 2008). The plasma membrane functions by creating a tight barrier between the cytoplasm and the exterior. This is achieved by a 3–5 nm hydrophobic barrier that has to be overcome by water soluble molecules before they enter or exit the cell (See > Chapter 29, Vol. 1, Part 5). Disruption of this barrier has grave consequences for most cells although their sensitivity varies (See > Chapter 43, Vol. 2, Part 7; > Chapter 44, Vol. 2, Part 7; > Chapter 52, Vol. 2, Part 9). Bacteria, with their small internal volumes, often unfriendly external environments and single membrane, which carries out many functions from nutrient uptake to energy conversion, are much more sensitive to small increases in permeability than eukaryotic cells. The latter are relatively large, in protected environments and possessors of a whole range of specialized membrane compartments (Iacovache et al., 2008; Lakey et al., 1994b). It is, however, possible to damage an animal cell which lacks the cell walls found in plants and fungi by colloid osmotic lysis. This occurs when the cell’s water permeability is increased by toxin induced pores. The cell is full of large macromolecules such as proteins, DNA etc which act as impermeable colloids which will not equilibrate across the bilayer. On the other hand water and small solutes enter the cell in response to a lowered water potential and concentration gradients respectively. The cell is unable to mitigate this flux and increases in volume until it lyses. Nevertheless, it has been found that membrane damaging proteins whose targets are eukaryotic often have subtle effects in intracellular compartments which do not rely upon the bursting of the cell (Iacovache et al., 2008). The clearest examples of these are the toxin delivering proteins which penetrate the target cell membrane in order to deliver cargo enzymes that target intracellular sites with extreme selectivity. Examples of these are diphtheria, anthrax, PMT, cholera and verotoxins. Even bacterial toxins which appear to rely upon pore formation alone have effects upon cells at concentrations which are considered sub-lytic and may rely on their effects upon intracellular compartments; these include aerolysin, alpha-hemolysin and the increasingly medically important LukF toxin from Staphylococcus aureus. In both groups, toxins have evolved to use intracellular compartments by receptor mediated endocytosis. Members of this group of toxin deliverers appear to cross the intracellular membranes efficiently and do not appear to cause observable damage to the lipid bilayer itself (Iacovache et al., 2008).

Membrane Disrupting Proteins

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This requirement for endocytosis reveals a feature of many membrane damaging proteins which is their reliance upon surface receptors. The toxins must have the ability to be both water and membrane soluble and have been termed Janus proteins after the Roman God who could look both into the past and the future and was often depicted upon doorways looking both outward and inside the house. To make the transition from water to the membrane, the proteins much have an affinity for some aspect of the membrane surface. The most obvious of these are proteinacious cell surface receptors first recognized by Ehrlich (1913). However many membrane damaging molecules have the ability to interact directly with the membrane bilayer itself. This review will cover the transition of membrane damaging proteins from aqueous to membrane state and address some basic features of this process.

2

The Importance of Secondary Structure

Insertion of proteins into membranes is dependent upon the efficient masking of the peptide bond. In every n-residue peptide/protein there are (n-1) peptide bonds (not taking into account the imide bonds of X-proline linkages) so they are both universal and are as numerically important as side chains. If alone they will expose both an amide proton and carbonyl oxygen which, because of the nature of the peptide bond, are highly polarized hydrogen bond donors and acceptors respectively. This makes them unfavorable membrane components unless they can replace water, as a hydrogen bonding partner, with something else. In fact. burial of a hydrogen bond in the membrane incurs an energy cost approximating to that of charged side chains. The most efficient way to ensure the neutralization of the peptide bond polarity is to form hydrogen bonds with other peptide bonds by the formation of an alpha helix or beta sheet structure (> Fig. 1). It is because of this that an understanding of the role of secondary structure in membrane damaging peptides and proteins is so crucial. In the majority of cases the proteins fall into one or other secondary structure family and this is understandable when their relative properties are understood.

3

The Alpha-Helix

Alpha helices, as will be exemplified by the bee venom peptide melittin, create a hydrogen bonding network that is contained within one contiguous section of polypeptide (> Fig. 1a). Thus if there are flexible links between the helices in large proteins these can act as separate structural units (> Fig. 1b) only interacting if they have complementary surfaces defined by their side chains, which always point outwards from the helix. The individual helices can be water soluble, hydrophobic or as with melittin amphipathic. In the case of poly-helical membrane damaging proteins the elements arrange themselves according to similar rules but in this case guided by the inter helix links (Kolter et al., 2005). The alpha helix thus represents the simplest form of protein secondary structure for insertion into membranes. Its hydrophobicity is determined by the side chains which point outwards from the helix and here the nature of the membrane interaction is defined. The example chosen here is melittin although there are many other examples of so called lytic peptides (Bechinger, 2004). Melittin is a 26 residue cationic peptide from the venom of the European honey bee (Apis mellifera) (> Fig. 2) (Raghuraman and Chattopadhyay, 2007). The helix forms an

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. Figure 1 Protein interactions with membranes. (a) Helices can adopt a surface bound arrangement or form more deeply inserting structures. This is regulated by helix hydrophobicity, density of peptides at the surface and lipid interactions. (b) The same general rules affect proteins made of groups of helices except that the helix-helix links restrict the rearrangement and because their peptide bonds may not be hydrogen bonded they may be unstable within the membrane core. (c) Beta structure forms hydrogen bonds between strands (dotted lines) but at the end of a beta sheet the structure cannot expose edges to the bilayer core. Thus beta barrels are the usual form (see > Fig. 3c).

amphipathic structure and can make a molecule with the required Janus behavior (> Fig. 2a, b). Every third to fourth residue will point in one direction and, if hydrophobic, the amphipathic helix motif can be observed readily even in the primary structure. In some cases, such as melittin, the peptide may be unfolded in solution and then adopt the helix only when bound to the membrane (Bechinger, 2004). In this way the water solubility of the peptide is high in water due to the free peptide bonds and then it can adopt the necessary hydrophobic helix in the membrane. The attraction to the membranes surface is thus an important step and the peptide must first be attracted to the hydrophobic membrane. In most cases the lytic peptides are cationic and since most membranes bear a net negative charge it is their lysine and arginine side chains that promote the first interactions. The long side chains of lysine and arginine also provide hydrophobic groups and thus play a critical amphiphilic role in the membrane interaction (> Fig. 2c). Cationic anti-microbial peptides are in fact an essential part of innate immunity in both plants and animals (Bechinger and Lohner, 2006). The other specific interaction is driven by tryptophan (see later). The cartoon representation of melittin (> Fig. 2d) reveals a break in the helix and this is due to a proline at position 14.

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. Figure 2 Details of melittin; an example of a cationic membrane damaging peptide. (a and b) Two different views of the helical form of melittin with the hydrophobic surfaces colored dark. (c) End view with the membrane interaction surface pointing downwards and key Trp and cationic side chains highlighted. (d) Cartoon of helix structure showing the break in the peptide caused by the Proline14 residue.

This creates a flexible backbone and is a feature of several lytic peptides. The lytic peptides have varied roles in membrane damage. In bee venom for example they induce phospholipase activity but more generally they can induce pores which can kill microbial and eukaryotic cells. There are several models of how the initial surface binding develops into pore formation. The fundamental two types are a transmembrane barrel (barrel stave or wormhole models) with their water soluble surfaces facing the pore lumen and a carpet/detergent model where the density of lytic peptides at the membrane surface causes a break in the membrane structure. The tight binding of peptides to lipids and the dynamic states have also suggested a toroidal pore in which the structure is a mixture of lipids and proteins (> Fig. 1a). The membrane disrupting proteins which are composed of helices (Iacovache et al., 2008; Kolter et al., 2005; Lakey et al., 1994b) comprise various combinations of hydrophobic, hydrophilic and amphipathic helices. They are folded in water solution with the hydrophobic sections buried (> Fig. 3b) and then upon membrane binding unfold to allow the hydrophobic components to insert into the membrane and form membrane disrupting structures. Usually the helix content is largely unchanged and a rearrangement is the main feature of the insertion step (Anderluh and Lakey, 2008; Lakey et al., 1994b).

4

Beta Structure

Beta-type membrane damaging toxins can be separated into those in which the beta structure remains outside of the membrane and those in which it forms the membrane penetrating structure. In the former, such as the example of equinatoxin from the sea anemone Actinia equina, the beta structure provides a rigid scaffold to which are attached lipid binding and membrane insertion structures (> Fig. 3a). In the later the beta domain penetrates the membrane and has to conform to the limitations of polypeptide in the membrane (> Fig. 3c). The nature of beta secondary structure is determined by hydrogen bonding

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. Figure 3 Structures of membrane damaging proteins. (a) Equinatoxin (PDB:1IAZ) has a rigid beta sandwich core which does not penetrate the membrane. A tryptophan rich loop (space filling representation) binds the membrane interface and contains a sphingomyelin pocket whilst the amphipathic (melittin like) N terminal helix (shaded dark; left) moves downwards into the membrane core to create a pore in an oligomer. (b) Pore-forming domain from colicin N, an antibacterial toxin from Escherichia coli (PDB:1A87). This is an example of a helical water soluble toxin that rearranges its structure to create a membrane spanning pore. The dark helices are fully hydrophobic and kept soluble by the amphipathic melittin like peptides around them. (c) The alpha hemolysin from Staphylococcus aureus pore is composed of seven monomers each of which contributes a single beta hairpin (highlighted dark) to the barrel that crosses the membrane (PDB:7AHL). The outside of the barrel exposes hydrophobic side chains whilst the alternate side chains are hydrophilic and point inwards making a non specific water filled pore.

between the peptide bonds of separate beta strands. The strands may be consecutive in the primary structure but in all cases at least a pair (minimally a beta hairpin) are needed to define the beta structure. Whereas the free edge of a beta sheet in soluble proteins can interact with water, upon membrane insertion the peptides need to make complete sets of hydrogen bonding interactions. This is illustrated in > Fig. 1c and it is clear that a flat beta sheet cannot be energetically stable in the hydrophobic core of the bilayer. The solution to this, found in all beta sheet membrane disrupting proteins, is the rolling up of the beta sheet into a beta barrel. Thus the free edges of the sheet become just another bonded strand-strand interface.

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The structure is the same as outer membrane proteins from bacteria and mitochondria but in general these barrels are composed of one polypeptide. In membrane damaging beta barrel toxins the water to membrane bound step usually involves an essential oligomerization and thus the membrane penetrating beta barrel is composed of beta hairpins from several monomers. The clearest examples are the heptameric pore-forming toxins such as alpha hemolysin (> Fig. 3). The 14-stranded barrel is formed by beta hairpins from seven monomers. These hairpins are often unrecognizable in the water soluble monomer which first binds to the surface via a receptor binding step followed by a tryptophan dependent interfacial interaction. Once in an oligomeric pre-pore state it develops into the membrane inserted form by extending the hydrophobic barrel into the bilayer. The side chains of a beta structure alternately point inwards and outwards down a strand and thus if every other side chain is hydrophobic it ensures that the final structure is membrane compatible. This also means that the unassembled hairpin is only semi hydrophobic and does not detract from the water solubility of the soluble monomer. The cholesterol dependent cytolysins produced by bacteria display a different arrangement in which the number of monomers is much greater (>30) and thus large ring shaped pores are formed in the target cell membrane.

5

Tryptophan

The role of tryptophan in the movement of peptides from water to membrane has proven to be surprisingly important. As it is the largest side chain and clearly hydrophobic, the likelihood that it would play a major role in membrane insertion could have easily been taken for granted. On closer inspection, however, it displays properties that have proven essential in the activity of several membrane damaging proteins. The replacement of the single tryptophan in melittin significantly reduces activity. In the actinoporin Equinatoxin II (> Fig. 3a), tryptophan residues are present on the loop which first interacts with the membrane and their replacement inhibits both binding to artificial membranes and in vivo toxicity (Hong et al., 2002). As in most biological interactions the on-rate for binding is diffusion limited and possibly affected in the short range by electrostatics (e.g., cationic peptides) which can accelerate the approach over the last few nm and even play a role in orienting the protein correctly at the membrane surface (Lakey et al., 1994a). Nevertheless it is the off rate that really determines the observed affinity and the insertion process requires an initial surface binding to provoke the structural changes that accompany the main insertion event. Specific side chains are thus fundamental for keeping the toxin at the membrane interface long enough for the second, largely irreversible stage of insertion to occur. Often the tryptophans provide a finger hold on the membrane by providing a way to catch hold of this generic interface. The next stages of EqtII membrane insertion are the specific binding to a sphingomyelin headgroup followed by the insertion of an amphipathic (melittin like) helix but this will not occur without the initial tryptophan interaction. The explanation for the behavior of tryptophan has largely been provided by the work of White and colleagues (Wimley and White, 1996; Yau et al., 1998) who developed more definitive versions of the original amino acid hydrophobicity scales. White and co introduced the interfacial scale. This was supported by measurements on standard peptides in which the central residue was changed for each of the 20 possible variants. The tryptophan behavior is thought to be due to pi electrons which favorably interact with the interface (Yau et al., 1998). On integral membrane proteins, tyrosines are also found with their hydroxyl groups pointing outwards across the interface but because this orientation makes

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them unlikely anchors for invading proteins it is tryptophan which is clearly the most important in interfacial binding. The Trp loop of EqtII is also rich in Tyr and Phe (Hong et al., 2002) and this pattern is also found in quite unrelated proteins such as the cholesterol dependent cytolysins and anthrax toxin (Anderluh and Lakey, 2008) and the sphingomyelinase from Listeria (Openshaw et al., 2005). In cholesterol dependent cytolysins the loop is well characterized by X-ray crystallography and has been studied in depth (Anderluh and Lakey, 2008). It is thought to be involved in the cholesterol binding and is another example of adopting an interfacial interaction in order to bind a poorly exposed membrane buried receptor.

6

Conclusion

Membrane damaging proteins travel to their target as water soluble precursors and must drive their own membrane binding and insertion steps. The critical phase of interaction is the interfacial binding and this is driven by cationic and tryptophan interactions. The subsequent step is reliant upon hiding the peptide bonds from the membrane core and this is achieved by the specific formation of secondary structure with hydrophobic side chains exposed in specific patterns.

7

Research Needs

In spite of high resolution X-ray structures for membrane damaging proteins in their water soluble state we have little idea how they form pores in the membrane. Methods that provide a clear picture of these dynamic structures in the membrane are constantly being sought but because of their dynamism these disrupted membranes are likely to keep their detailed functions a secret for a while yet.

References Anderluh G, Lakey JH (2008) Disparate proteins use similar architectures to damage membranes. Trends Biochem Sci 33: 482–490. Bechinger B (2004) Structure and function of membrane-lytic peptides. Crit Rev Plant Sci 23: 271–292. Bechinger B, Lohner K (2006) Detergent-like actions of linear amphipathic cationic antimicrobial peptides. Biochim Biophys Acta 1758: 1529–1539. Ehrlich P (1913) Address in pathology chemotherapeutics: scientific principle, methods and results. Lancet 182: 445–451. Hong Q, Gutierrez-Aguirre I, Barlic A, Malovrh P, Kristan K, Podlesek V, Macek P, GonzalezMan˜as J-M, Lakey JH, Anderluh G (2002) Two-step membrane binding by Equinatoxin II, a pore-forming toxin from the sea anemone. Involves

an exposed aromatic cluster and a flexible helix. J Biol Chem 277: 41916–41924. Iacovache I, van der Goot FG, Pernot L (2008) Pore formation: an ancient yet complex form of attack. Biochim Biophys Acta 1778: 1611–1623. Kolter T, Winau F, Schaible UE, Leippe M, Sandhoff K (2005) Lipid-binding proteins in membrane digestion, antigen presentation, and antimicrobial defense. J Biol Chem 280: 41125–41128. Lakey JH, Parker MW, Gonzalez Manas JM, Duche D, Vriend G, Baty D, Pattus F (1994a) The role of electrostatic charge in the membrane insertion of colicin A: calculation and mutation. Eur J Biochem 220: 155–163. Lakey JH, van der Goot FG, Pattus F (1994b) All in the family: the toxic activity of colicins. Toxicology 87: 85–108.

Membrane Disrupting Proteins Openshaw AEA, Race PR, Monzo HJ, Vazquez-Boland JA Banfield MJ (2005) Crystal structure of SmcL, a bacterial neutral sphingomyelinase C from Listeria. J Biol Chem 280: 35011–35017. Raghuraman H, Chattopadhyay A (2007) Melittin: a membrane-active peptide with diverse functions. Biosci Rep 27: 189–223.

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Wimley WC, White SH (1996) Experimentally determined hydrophobicity scale for proteins at membrane interfaces. Nat Struct Biol 3: 842–848. Yau WM, Wimley WC, Gawrisch K White SH (1998) The preference of tryptophan for membrane interfaces. Biochemistry 37: 14713–14718.

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31 Lipid Intermediates in Bacterial Peptidoglycan Biosynthesis J. van Heijenoort Institute of Biochemistry and Molecular and Cellular Biophysics, University Paris-Sud, Orsay, France [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 436 2 Lipid Intermediates in the Biosynthesis of Peptidoglycan . . . . . . . . . . . . . . . . . . . . . . . . . . 436 3 Biosynthesis of Lipid I by Transferase MraY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 437 4 Biosynthesis of Lipid II by Transferase MurG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 439 5 Biosynthesis of Modified Lipid Intermediates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 440 6 Cellular Location and Translocation of Lipid Intermediates . . . . . . . . . . . . . . . . . . . . . . . 441 7 Glycan Chain Formation by Transglycosylation with Lipid II . . . . . . . . . . . . . . . . . . . . . . 441 8 Complexes Between the Lipid Intermediates and Antibiotics . . . . . . . . . . . . . . . . . . . . . . . 442 9 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 443

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Abstract: Lipid intermediates I and II involved in the biosynthesis of bacterial peptidoglycan are the undecaprenyl pyrophosphoryl MurNAc-pentapeptide and undecaprenyl pyrophosphoryl GlcNAc-b-(1!4)-MurNAc-disaccharide-peptide, respectively. Membrane transferase MraY catalyzes the formation of lipid I whereas peripheral MurG transferase catalyzes the addition of N-acetylglucosamine onto lipid I to yield lipid II. These lipid intermediates often undergo additional modifications leading to complex pools. The glycan chains of peptidoglycan are assembled by polymerization of the GlcNAc-b-(1!4)-MurNAc-disaccharide-peptide unit of lipid II with formation of b-(1!4) linkages. Antibiotics such as glycopeptides and lantibiotics bind noncovalently with extracytoplasmically located lipid II and thereby lead to the arrest of peptidoglycan polymerization. Chemically or enzymatically synthesized lipid intermediates and analogues are now available in useful amounts for the study of the membrane steps of peptidoglycan biosynthesis, of the complexes formed with antibiotics, and of mechanisms of antibiotic resistance. Future work will require the development of convenient methods for the analysis of the lipid pools. Among the cellular aspects still poorly addressed, the elucidation of the mechanism of translocation of lipid II through the membrane remains an important challenging enigma.

1

Introduction

Bacterial peptidoglycan has been extensively investigated owing to its importance as an essential structural cell wall component (Vollmer et al., 2008), to its involvement in cellular morphogenesis (den Blaauwen et al., 2008; Zapun et al., 2008), and to the fact that steps of its biosynthesis are specifically inhibited by well-known antibiotics and are potential targets for the search of novel antibacterials (Barreteau et al., 2008; Gale et al., 1981; Green, 2002; Kotnik et al., 2007). Its biosynthesis is a two-stage process. First, the peptidoglycan monomer unit is assembled (> Fig. 1) in the final form of a lipid intermediate by enzymes located in the cytoplasm or at the inner side of the cytoplasmic membrane (Barreteau et al., 2008; Bouhss et al., 2008; van Heijenoort, 2001b, 2007). In the second stage, extracytoplasmic glycosyltransferases catalyze the formation of linear glycan chains using the last lipid intermediate as substrate (Goffin and Ghuysen, 1998; van Heijenoort, 2001a, 2007). Finally, the cross-linking between the peptide subunits by DD-transpeptidases leads to the 3D network of peptidoglycan (Goffin and Ghuysen, 2002; Macheboeuf et al., 2006; Sauvage et al., 2008).

2

Lipid Intermediates in the Biosynthesis of Peptidoglycan

The study of cell-free peptidoglycan-synthesizing systems using UDP-GlcNAc, radiolabeled UDP-MurNAc-pentapeptide (> Fig. 1) and bacterial membranes led to the discovery of lipid intermediates I and II, to the determination of their role in the pathway, and to that of their structures (> Fig. 2) as undecaprenyl pyrophosphoryl MurNAc-pentapeptide and undecaprenyl pyrophosphoryl GlcNAc-b-(1!4)-MurNAc-disaccharide-peptide, respectively (Bouhss et al., 2008; van Heijenoort, 2007). The first membrane step (> Fig. 2) involves the transfer of the phospho-MurNAc-pentapeptide moiety of UDP-MurNAc-pentapeptide to undecaprenyl phosphate to yield lipid I and UMP. Thereafter, N-acetylglucosamine is added onto lipid I to yield lipid II. The pool levels of the lipid intermediates are very low in comparison with the cell lipid content. In E. coli, there are at best a few thousand copies per cell whereas in Grampositive bacteria estimates range from 34,000 to 200,000 copies per cell (van Heijenoort, 2007).

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. Figure 1 Stepwise assembly of the peptidoglycan monomer unit. GlcNAc N-acetylglucosamine, MurNAc N-acetylmuramic acid, A2pm diaminopimelic acid.

Their low pool levels, their limited accumulation in cell-free systems, and the tedious work of their isolation restricted for decades their availability for the study of the membrane steps of the pathway. These difficulties have now been overcome by their chemical or enzymatic synthesis (Bouhss et al., 2008; van Heijenoort, 2007; Welzel, 2005).

3

Biosynthesis of Lipid I by Transferase MraY

The formation of lipid I from UDP-MurNAc-pentapeptide and undecaprenyl phosphate is catalyzed by integral membrane protein MraY which is essential and unique (Boyle and

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. Figure 2 Structure of lipids I and II, and their synthesis by transferases MraY and MurG. Pep pentapeptide.

Donachie, 1998; Ikeda et al., 1991). MraY belongs to the UDP-D-N-acetylhexosamine: polyprenol phosphate D-N-acetylhexosamine 1-P transferase family (Price and Momany, 2005). MraY has been purified but not yet crystallized (Bouhss et al., 2004). The 2D membrane topology model of MraY shows that it has ten transmembrane segments, five cytoplasmic loops and six periplasmic loops including the N- and C-terminal ends. The cytoplasmic loops are involved in substrate recognition and catalysis, thereby indicating that lipid I is formed at the inside surface of the membrane (Bouhss et al., 1999; Price, and Momany, 2005). A multistep catalytic mechanism (> Fig. 3) was proposed (van Heijenoort, 2007). The MraY reaction is fully reversible and takes place with conservation of the a-anomeric configuration

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. Figure 3 Multistep MraY-catalyzed formation of lipid I. R D-lactoyl-peptide.

of the MurNAc residue. Many inhibitors of MraY have been described (Dini, 2005). Furthermore, it is noteworthy that undecaprenyl phosphate is also involved in the biosynthesis of various other cell-wall components (Bouhss et al., 2008).

4

Biosynthesis of Lipid II by Transferase MurG

The transfer of N-acetylglucosamine from UDP-GlcNAc onto the C4 hydroxyl of the MurNAc unit of lipid I is catalyzed by transferase MurG (Mengin-Lecreulx et al., 1991). The formation of the b-(1!4) linkage is accompanied by inversion of the anomeric configuration of

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. Figure 4 MurG-catalyzed formation of lipid II. R D-lactoyl-peptide.

N-acetylglucosamine (> Fig. 4). MurG belongs to the GT-B glycosyltransferase superfamily ¨ nligil and Rini, 2000). It is unique, essential, and shown to be associated with the inner side (U of the cytoplasmic membrane thereby establishing that the entire peptidoglycan monomer unit is assembled inside the cell prior to translocation across the membrane. MurG is thus a key enzyme at the junction between the two stages of peptidoglycan synthesis (Bouhss et al., 2008; van Heijenoort, 2007). In the 3D structure of MurG the binding site for UDP-GlcNAc is in the C-terminal domain and that for lipid I in the N-terminal domain, in which a hydrophobic patch surrounded by basic residues is perhaps the site of interaction with the negatively charged membrane (Ha et al., 2000). Sequence comparison of various orthologues confirmed the extrinsic and cationic character of MurG, and its functioning as a moderately hydrophobic peripheral protein. Detailed enzymatic studies were carried out with lipid I analogues (Bouhss et al., 2008; van Heijenoort, 2007).

5

Biosynthesis of Modified Lipid Intermediates

Variations in the basic structure of peptidoglycan are encountered in most bacteria and are introduced at different steps of the biosynthesis (Vollmer et al., 2008). Those undergone by the lipid intermediates concern amidation or esterification of the peptide subunit carboxyls or addition of extra amino acids leading, in particular, to the presence of peptide bridges between the peptide subunits in many Gram-positive bacteria (van Heijenoort, 2007). Another

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important modification is the formation of a protein-associated lipid II intermediate during the cell wall sorting pathway of surface proteins (Marraffini et al., 2006). Since the various modifications are not necessarily complete, lipid intermediates are in many cases complex mixtures of modified forms.

6

Cellular Location and Translocation of Lipid Intermediates

Since lipid II (or a modified form) is entirely assembled on the inner surface of the cytoplasmic membrane, it must be translocated through the hydrophobic environment of the membrane prior to its use as substrate in extracytoplasmic polymerization (Bouhss et al., 2008; van Heijenoort, 2001a, 2007). An E. coli growing cell has ca. 2,000 copies of lipid II and peptidoglycan polymerization proceeds at ca. 1,000 units per second. This implies a turnover of a few seconds for the lipid II pool. The translocation of lipid II must be a fast unidirectional process in order to sustain a steady peptidoglycan synthesis. More than four decades after its discovery, the mechanism of this translocation remains unknown. Fluorescence spectrometry experiments carried out in lipid vesicles showed that there was no spontaneous move of the lipid intermediate across the bilayer (van Dam et al., 2007). Nevertheless, translocation was observed when E. coli inner membrane vesicles were used, revealing that a translocationmachinery was present and likely composed of membrane proteins. Essential inner-membrane protein Mur J (or MviN) of E. coli has just been identified as the likely lipid II translocase (Ruiz, 2008; Inoue et al., 2008).

7

Glycan Chain Formation by Transglycosylation with Lipid II

The glycan chains of peptidoglycan are assembled by polymerization of the GlcNAc-b-(1!4)MurNAc-disaccharide-peptide unit of lipid II with formation of b-(1!4) linkages (> Fig. 5). The transglycosylation reaction is accompanied by inversion of the a-anomeric configuration of the MurNAc residue and thus leads to linear chains containing exclusively b-(1!4) linkages (van Heijenoort, 2001a). In several cases, the reaction was shown to occur at the reducing end of the growing chain which is transferred as the donor substrate to the C4 hydroxyl of the GlcNAc unit of lipid II acting as the acceptor substrate (Perlstein et al., 2007; van Heijenoort, 2001a). Undecaprenyl pyrophosphate, which is the side product of the polymerization reaction (> Fig. 5) and the target of bacitracin, is recycled back into the cell. Together with undecaprenyl pyrophosphate synthesized endogenously by the UppS synthase, it is dephosphorylated by membrane phosphatases to undecaprenyl phosphate which is available for the MraY reaction and the synthesis of other cell wall components (Bouhss et al., 2008). The glycosyltransferases (GTs) responsible for the polymerization reaction come in two forms (Goffin and Ghuysen, 1998; van Heijenoort, 2001a, 2007), namely an N-terminal module in bifunctional class A penicillin-binding proteins, which also contain a C-terminal transpeptidase module, and monofunctional glycosyltransferases. They show high sequence similarity, belong to the GT51 family in the sequence-based classification of glycosyltransferases, and possess five conserved motifs (Coutinho et al., 2003; Goffin and Ghuysen, 1998). Presently, over 15 peptidoglycan GTs have been purified, and most assayed for their activity with lipid II (Sauvage et al., 2008; van Heijenoort, 2007). Analysis of the reaction products formed by purified E. coli GT PBP1b with lipid II revealed an average glycan chain length of over 25 disaccharide units and 50% of cross-linked

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. Figure 5 Mechanism of transglycosylation with chain elongation at the reducing end. R D-lactoyl-peptide, GT glycosyltransferase.

peptides (Bertsche et al., 2005). The recent resolution of the crystal structures of PBP2 from Staphylococcus aureus (Lovering et al., 2007) and of the GT domain of Aquifex aeolicus (Yuan et al., 2007) has opened the way to more systematic studies of the GT catalytic site and reaction mechanism.

8

Complexes Between the Lipid Intermediates and Antibiotics

Glycopeptides (vancomycin, teicoplanin, telavancin, . . .), lantibiotics (nisin, mersacidin, epidermin, . . .), and other antibiotics (ramoplanin, mannopeptide, . . .) can bind noncovalently to the peptidoglycan lipid intermediates (Bauer and Dicks, 2005; Breukink and de Kruijff, 2006; Chatterjee et al., 2005; Kahne et al., 2005; Walker et al., 2005). In general, they are cyclic peptides, depsipeptides or peptides with extra posttranslationally introduced ring rearrangements and amino acid modifications. Some contain sugar and/or lipid moieties. Owing to their size and polar structure, they penetrate with difficulty both the bacterial cytoplasmic membrane and the outer membrane of Gram-negative bacteria. Therefore, in Gram-positive bacteria they interact with extracytoplasmically located lipid II and by its sequestration they lead to the arrest of peptidoglycan polymerization. However, these antibiotics differ from one another in many respects such as their affinity for lipid II, the interacting sites of lipid II, and the possession of additional and separate mechanisms of action affecting their antibacterial activity. Lipid II-antibiotic complexes are used for the study of the dynamic assembly of peptidoglycan (Tiyanont et al., 2006).

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Research Needs

Chemically or enzymatically synthesized lipid intermediates and analogues are now available in useful amounts for the study of the membrane steps of peptidoglycan biosynthesis, of the complexes formed with many antibiotics, and of mechanisms of antibiotic resistances. Various modifications of lipids I and II have been described and undoubtedly more are yet to be identified. They often lead to complex pools the analysis of which is difficult but essential for understanding the in vivo functioning of the membrane steps and that of possible regulatory mechanisms. Future work will require the development of convenient and efficient analytical methods. Moreover, the biochemistry of the reactions responsible of modifications has been studied only in a few cases and much remains to be done. Among the cellular aspects still poorly addressed, the elucidation of the mechanism of translocation of lipid II or its modified forms remains an important challenging problem. This will require the development of appropriate translocation assays to be used for the systematic search of the membrane proteins involved in the process.

References Barreteau H, Kovacˇ A, Boniface A, Sova M, Gobec S, Blanot D (2008) Cytoplasmic steps of peptidoglycan synthesis. FEMS Microbiol Rev 32: 168–207. Bauer R, Dicks LMT (2005) Mode of action of lipid II-targeting lantibiotics. Int J Food Microbiol 101: 201–216. Bertsche U, Breukink E, Kast T, Vollmer W (2005) In vitro murein (peptidoglycan) synthesis by dimers of the bifunctional transglycosylase-transpeptidase PBP1B from Escherichia coli. J Biol Chem 280: 38096–38101. Bouhss A, Crouvoisier M, Blanot D, Mengin-Lecreulx D (2004) Purification and characterization of the bacterial MraY translocase catalyzing the first membrane step of peptidoglycan biosynthesis. J Biol Chem 279: 29974–29980. Bouhss A, Mengin-Lecreulx D, Le Beller D, van Heijenoort J (1999) Topological analysis of the MraY protein catalysing the first membrane step of peptidoglycan synthesis. Mol Microbiol 34: 576–585. Bouhss A, Trunkfield AE, Bugg TDH, Mengin-Lecreulx D (2008) The biosynthesis of peptidoglycan lipidlinked intermediates. FEMS Microbiol Rev 32: 208–233. Boyle DS, Donachie WD (1998) mraY is an essential gene for cell growth in Escherichia coli. J Bacteriol 180: 6429–6432. Breukink EI, de Kruijff B (2006) Lipid II as a target for antibiotics. Nat Rev Drug Discov 4: 321–332. Chatterjee C, Paul M, Xie L, van der Donk WA (2005) Biosynthesis and mode of action of lantibiotics. Chem Rev 105: 633–683.

Coutinho PM, Deleury E, Davies GJ, Henrissat B (2003) An evolving hierarchical family classification for glycosyltransferases. J Mol Biol 328: 307–317. den Blaauwen T, de Pedro MA, Nguyen-Diste`che M, Ayala JA (2008) Morphogenesis of rod-shaped sacculi. FEMS Microbiol Rev 32: 321–344. Dini C (2005) MraY inhibitors as novel antibacterial agents. Curr Top Med Chem 5: 1221–1236. Gale EF, Cundiffe E, Reynolds PE, Richmond MH, Warning MJ (1981) The Molecular Basis of Antibiotic Action. London: John Wiley and Sons. Goffin C, Ghuysen J-M (1998) Multimodular penicillinbinding proteins: an enigmatic family of orthologs and paralogs. Microbiol Mol Biol Rev 62: 1079–1093. Goffin C, Ghuysen J-M (2002) Biochemistry and comparative genomics of SxxK superfamily acyltransferases offer a clue to the mycobacterial paradox: presence of penicillin-susceptible target proteins versus lack of efficiency of penicillin as therapeutic agent. Microbiol Mol Biol Rev 66: 702–738. Green DW (2002) The bacterial cell wall as a source of antibacterial targets. Expert Opin Ther Targets 6: 1–19. Ha S, Walker D, Shi Y, Walker S (2000) The 1.9 A˚ crystal structure of Escherichia coli MurG, a membraneassociated glycosyltransferase involved in peptidoglycan biosynthesis. Protein Sci 9: 1045I–1052I. Ikeda M, Wachi M, Jung HK, Ishino F, Matsuhashi M (1991) The Escherichia coli mraY gene encoding UDPN-acetylmuramoyl-pentapeptide: undecaprenylphosphate phospho-N-acetylmuramoyl-pentapeptide transferase. J Bacteriol 173: 1021–1026.

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Inoue A, Murata Y, Takahashi H, Tsuji N, Fujisaki S, Kato J (2008) Involement of an essential gene, mviN, in murein synthesis in Escherichia coli. J Bacteriol 190: 7298–7301. Kahne D, Leimkuhler C, Lu W, Walsh C (2005) Glycopeptide and lipoglycopeptide antibiotics. Chem Rev 105: 425–448. Kotnik M, Anderluh PS, Prezelj A (2007) Development of novel inhibitors targeting intracellular steps of peptidoglycan biosynthesis. Curr Pharma Design 13: 2283–2309. Lovering AL, de Castro LH, Lim D, Strynadka NCJ (2007) Structural insight into the transglycosylation step of bacterial cell-wall biosynthesis. Science 315: 1402–1405. Macheboeuf P, Contreras-Martel C, Job V, Dideberg O, Dessen A (2006) Penicillin binding proteins: key players in bacterial cell cycle and drug resistance processes. FEMS Microbiol Rev 30: 673–691. Marraffini LA, Dedent AC, Schneewind O (2006) Sortases and the art of anchoring proteins to the envelopes of gram-positive bacteria. Microbiol Mol Biol Rev 70: 192–221. Mengin-Lecreulx D, Texier L, Rousseau M, van Heijenoort J (1991) The murG gene of Escherichia coli codes for the UDP-N-acetylglucosamine:Nacetylmuramyl-(pentapeptide) pyrophosphorylundecaprenol N-acetylglucosamine transferase involved in the membrane steps of peptidoglycan synthesis. J Bacteriol 173: 4625–4636. Perlstein DL, Zhang Y, Wang T-S, Kahne DE, Walker S (2007) The direction of glycan chain elongation by peptidoglycan glycosyltransferases. J Am Chem Soc 129: 12674–12675. Price NP, Momany FA (2005) Modeling bacterial UDPhexNAc:polyprenol-P hexNAc-1-P transferases. Glycobiology 15: 29R–42R. Ruiz N (2008) Bioinformatics identification of MurJ (MviN) as the peptidoglycan lipid II flippase in Escherichia coli. Proc Natl Acad Sci USA 105: 15553–15557.

Sauvage E, Kerff F, Terrak M, Ayala JA, Charlier P (2008) The penicillin-binding proteins: structure and role in peptidoglycan synthesis. FEMS Microbiol Rev 32: 234–258. Tiyanont K, Doan T, Lazarus MB, Fang X, Rudner DZ, Walker S (2006) Imaging peptidoglycan biosynthesis in Bacillus subtilis with fluorescent antibiotics. Proc Natl Acad Sci USA 103: 11033–11038. ¨ nligil UM, Rini JM (2000) Glycosyltransferase structure U and mechanism. Curr Opin Struct Biol 10: 510–517. van Dam V, Sijbrandi R, Kol M, Swiezewska E, de Kruijff1 B, Breukink E (2007) Transmembrane transport of peptidoglycan precursors across model and bacterial membranes. Mol Microbiol 64: 1105–1114. van Heijenoort J (2001a) Formation of the glycan chains in the synthesis of bacterial peptidoglycan. Glycobiology 11: 25R–36R. van Heijenoort J (2001b) Recent advances in the formation of the bacterial peptidoglycan monomer unit. Nat Prod Rep 18: 503–519. van Heijenoort J (2007) Lipid intermediates in the biosynthesis of bacterial peptidoglycan. Microbiol Mol Bio Rev 71: 620–635. Vollmer W, Blanot D, de Pedro MA (2008) Peptidoglycan structure and architecture. FEMS Microbiol Rev 32: 149–167. Walker S, Chen L, Hu Y, Rew Y, Shin D, Boger DL (2005) Chemistry and biology of ramoplanin: a lipoglycodepsipeptide with potent antibiotic activity. Chem Rev 105: 449–475. Welzel P (2005) Syntheses around the transglycosylation step in peptidoglycan biosynthesis. Chem Rev 105: 4610–4660. Yuan Y, Barrett D, Zhang Y, Kahne D, Sliz P, Walker S (2007) Crystal structure of a peptidoglycan glycosyltransferase suggests a model for processive glycan chain synthesis. Proc Natl Acad Sci USA 104: 5348–5353. Zapun AT, Vernet T, Pinho MG (2008) The different shapes of cocci. FEMS Microbiol Rev 32: 345–360.

32 Phenolic Lipids Synthesized by Type III Polyketide Synthases A. Miyanaga . S. Horinouchi* Department of Biotechnology, Graduate School of Agriculture and Life Sciences, University of Tokyo, Tokyo, Japan *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 446 2 Biosynthesis of Phenolic Lipids in Azotobacter vinelandii . . . . . . . . . . . . . . . . . . . . . . . . . . 447 3 Biosynthesis of Phenolic Lipids in Streptomyces griseus . . . . . . . . . . . . . . . . . . . . . . . . . . . . 448 4 Allelochemical Biosynthesis in Sorghum bicolor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 449 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 449

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_32, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Phenolic lipids, consisting of polar aromatic rings and hydrophobic alkyl chains, are distributed widely in bacteria, fungi, and plants and appear to play an important role in the biologic membranes. Biosynthesis of such phenolic lipids in two bacteria, Azotobacter vinelandii and Streptomyces griseus, and one plant, Sorghum bicolor is summarized. The phenolic lipids are biosynthesized by a combination of fatty acid synthases (FASs) responsible for the synthesis of the alkyl chains and type III polyketide synthases responsible for the synthesis of the aromatic rings.

1

Introduction

Phenolic lipids, consisting of polar aromatic rings and hydrophobic alkyl chains, are distributed widely in bacteria, fungi, and plants (Kozubek and Tyman, 1999). The amphiphilic nature of phenolic lipids contributes to the formation of stable monomolecular layers in vitro. Phenolic lipids also exhibit antimicrobial and antioxidation activities. Gellerman et al. (1976) confirmed that a polyketide synthase (PKS) is involved in the biosynthesis of the aromatic ring moiety of phenolic lipids, by showing that 14C-labeled acetate is efficiently incorporated into the phenolic ring of anacardic acid in plant Ginkgo biloba. Moreover, recent studies (Dayan et al., 2003; Funa et al., 2006; Funabashi et al., 2008; Miyanaga et al., 2008) showed that the biosynthesis of phenolic lipids involves the convergence of the fatty acid and polyketide pathways. FASs and PKSs share a similar mechanism for the synthesis of fatty acids and polyketides, respectively. They possess a ketosynthase activity that catalyzes the sequential decarboxylative addition of some acetate units from extender substrates, such as malonyl-CoA, to an acyl starter substrate. Their substrates and intermediate products are maintained as thioester conjugates to acyl carrier protein (ACP) or small molecule coenzyme A (CoA). However, their reactions are different. FAS systems perform the reduction and dehydration reactions on each resulting b-keto carbon to produce an alkyl chain, whereas PKS systems omit some of reduction and dehydration reactions and generate monocyclic or polycyclic products, accompanied by intramolecular cyclization of the polyketide chain. FASs are classified into two groups. Type I FASs are large multifunctional enzymes that have a set of distinct catalytic domains and are distributed in mammals, fungi, and yeasts (Schweizer and Hofmann, 2004). In bacteria, however, the de novo synthesis of fatty acids is catalyzed by type II FASs, which are a group of monofunctional proteins with distinct properties (Rock and Cronan, 1996). The mechanism of termination of the reaction differs in the different types of FASs. Mammalian type I FASs hydrolyze acyl-ACP to yield a free acid by thioesterases (TEs), and fungal and yeast type I FASs transfer the acyl moiety to CoA by malonyl/palmitoyl transferases (MPTs). In contrast, type II FASs release their products as ACP esters without cleaving the thioester bond. PKSs fall into three groups. Type I PKSs are mutifunctional enzymes that are organized into modules, each of which harbors a set of distinct catalytic domains. Type II PKSs consist of a complex of subunits such as a ketosynthase and a chain length factor. Type III PKSs distributed in plants, fungi, and bacteria have the simplest architecture, a homodimer of ketosynthase (Austin and Noel, 2003). They have been thought to mainly use an acyl-CoA thioester as the starter substrate. They use the same active site for both condensation and cyclization reactions. In this review, recent studies of the biosynthetic pathways of phenolic lipids in Azotobacter vinelandii, Streptomyces griseus, and Sorghum bicolor are described.

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Biosynthesis of Phenolic Lipids in Azotobacter vinelandii

A. vinelandii is a Gram-negative, nitrogen-fixing soil bacterium that differentiates into metabolically dormant cysts under adverse environmental conditions (Lin and Sadoff, 1968). Phenolic lipids, such as alkylresorcinols and alkylpyrones (> Fig. 1a), are the major lipids in the cyst membrane (Reusch and Sadoff, 1983). The phenolic lipids presumably allow the cysts to resist desiccation in A. vinelandii. These phenolic lipids are essential for formation of the cyst membrane and to be synthesized by an ars operon (Funa et al., 2006). This operon consists of two type III PKS genes, arsB and arsC, and two type I FAS genes, arsA and arsD. Two type III PKS genes, arsB and arsC, were cloned and characterized (Funa et al., 2006). ArsB and ArsC exhibit different catalytic properties, although they share 71% amino acid sequence identity. ArsB synthesizes alkylresorcinol by catalyzing three condensation of malonyl-CoA with a long acyl starter substrate, whereas ArsC synthesizes alkylpyrone by catalyzing two or three condensation of malonyl-CoA with a long acyl starter substrate (> Fig. 1a). ArsB and ArsC yield the same tetraketide intermediate. The different catalytic properties between ArsB and ArsC depend on the mechanism of cyclization. ArsB catalyzes

. Figure 1 Proposed pathways for the biosynthesis of phenolic lipids in A. vinelandii (a), S. griseus (b), and S. bicolor (c).

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intramolecular C2-to-C7 (numbering from the cysteine thioester) aldol condensation, accompanied by thioester cleavage and decarboxylative loss of the C1 carbon as CO2, to yield alkylresorcinol. On the other hand, ArsC catalyzes intramolecular C5 oxygen-to-C1 lactonization to yield alkyltetraketidepyrone. ArsC also use the triketide intermediate to yield alkyltriketidepyrone. arsA and arsD were also cloned and characterized (Miyanaga et al., 2008). ArsA and ArsD are unique type I FASs because of their domain organizations distinct from those of known type I FASs and the lack of TE and MPT domains in them. In vitro study (Miyanaga et al., 2008) showed that ArsA and ArsD catalyze the synthesis of C22, C24, and C26 fatty acids solely from malonyl-CoA. Consistent with the lack of TE and MPT domains in ArsA and ArsD, the products of ArsA and ArsD remain attached to ArsA, probably the ACP domain of ArsA. The analysis of the localization of radioactivity derived from [14C]-malonyl-CoA in the sequential reaction by ArsA, ArsB, ArsC, and ArsD demonstrated that the fatty acids produced by ArsA and ArsD are directly transferred to type III PKSs, ArsB, and ArsC, where they serve as starter substrates for phenolic lipid synthesis (> Fig. 1a). This is the first demonstration that a type I FAS interacts directly with a type III PKS through substrate transfer. Although ArsB and ArsC can accept acyl-CoA as a starter substrate, the direct transfer system implies that their primary substrate is acyl-ACP. Recently, enzymatic reactions other than thioesterification or transesterification between ACP and CoA to release fatty acids from type I FASs were found in some organisms. For example, Pap5A catalyzes the transfer of mycocerosic acid analogs to an alcohol, phthiocerol, from the ACP domain of a type I FAS, Mas, in Mycobacterium tuberculosis (Trivedi et al., 2005). In addition, direct transfer of C6 fatty acid is predicted to occur between HexAB (type I FASs) and PksA (a type I PKS) in Aspergillus parasiticus (Watanabe and Townsend, 2002). It is also hypothesized that steely, a fusion protein of a type I FAS and a type III PKS, directly transfers acyl products attached to the ACP domain of the type I FAS to the active site of the type III PKS in Dictyostelium discoideum (Austin et al., 2006). Such a direct transfer presumably facilitates an efficient catalysis by a following enzyme.

3

Biosynthesis of Phenolic Lipids in Streptomyces griseus

In S. griseus, phenolic lipids are perhaps associated with the cytoplasmic membrane and confer resistance to b-lactam antibiotics on the host. The phenolic lipids in S. griseus are shown to be synthesized by an srs operon (Funabashi et al., 2008). The fatty acids of Streptomyces are biosynthesized from amino acid degradation products methylbutyryl- and isobutyryl-CoA as starter units by type II FAS and therefore consist primarily of branched chain fatty acids (Wallace et al., 1995). SrsA, a type III PKS, condenses one methylmalonyl-CoA and two malonyl-CoA with this branched chain acyl substrate to yield alkylresorcinols (> Fig. 1b). A unique catalytic property of SrsA is that it synthesizes alkylresorcinols with an aromatic ring having a C-methyl group that is derived from methylmalonyl-CoA as an extender substrate. The order of condensation of methylmalonyl- and malonyl-CoA is strictly regulated in the order of malonyl-CoA, malonyl-CoA, and methylmalonyl-CoA. The cyclization of SrsA occurs as an intramolecular C2-to-C7 aldol condensation of a tetraketide, as is for ArsB. The alkylresorcinols synthesized by SrsA are efficiently modified by the post-polyketide modification enzymes, SrsB and SrsC. SrsB acts as a methyltransferase to catalyze regiospecific methylation of the phenol group of alkylresorcinols, yielding alkylresorcinol methyl esters. SrsC act as a flavoprotein hydroxylase to catalyze regiospecific hydroxylation of the

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alkylresorcinol methyl ethers, followed by nonenzymatic oxidation of the unstable hydroquinones, resulting in the formation of alkylquinones.

4

Allelochemical Biosynthesis in Sorghum bicolor

Sorgoleone, produced in root hair cells of sorghum (Sorghum bicolor), is probably responsible for much of the allelopathic properties of sorghum root exudates against broadleaf and grass weeds. The biosynthesis of sorgoleone is initiated by fatty acid desaturases such as SbDES2 and SbDES3 (> Fig. 1c) (Pan et al., 2007). They consecutively synthesize an unusual 16:3 fatty acid possessing a terminal double bond from palmitoyl-CoA, which is contained predominantly in all sorghum tissues. Dayan et al. (2003) showed by labeling study that a PKS is involved in the biosynthesis of the quinone moiety of sorgoleone and detected PKS activity responsible for sorgoleone biosynthesis in a sorghum root hair extract. A type III PKS thus uses a 16:3 fatty acyl-CoA starter unit and produces a pentadecatrienyl resorcinol (alkylresorcinol) intermediate, although it has not yet been identified in S. bicolor. Recently, the presence of type III PKS was predicted by an expressed sequence tag (EST) analysis (Baerson et al., 2008). This alkylresorcinol is then methylated by an S-adenosylmethionine-dependent O-methyltransferase (SbOMT3) and subsequently dihydroxylated by a hydroxylase (possibly a P450 monooxygenase), yielding a hydroquinone form of sorgoleone (Baerson et al., 2008). Such a post-modification system is similar to that of the phenolic lipids synthesis by the srs operon.

5

Research Needs

Phenolic lipids, which are distributed widely in bacteria, fungi, and plants, are attractive targets for chemical and biological studies because of their various biological activities. The recent studies have revealed that type III PKSs play an important role in the biosynthesis of phenolic lipids. These type III PKSs are coupled to FASs, forming fascinating and complicated biosynthetic reactions. The understandings of their biosynthesis and their biological activities will be potent in practical applications.

References Austin MB, Noel JP (2003) The chalcone synthase superfamily of type III polyketide synthases. Nat Prod Rep 20:79–110. Austin MB, Saito T, Bowman ME, Haydock S, Kato A, Moore BS, Kay RR, Noel JP (2006) Biosynthesis of Dictyostelium discoideum differentiation-inducing factor by a hybrid type I fatty acid-type III polyketide synthase. Nat Chem Biol 2:494–502. Baerson SR, Dayan FE, Rimando AM, Nanayakkara NP, Liu CJ, Schro¨der J, Fishbein M, Pan Z, Kagan IA, Pratt LH, Cordonnier-Pratt MM, Duke SO (2008) A functional genomics investigation of allelochemical

biosynthesis in Sorghum bicolor root hairs. J Biol Chem 283:3231–3247. Dayan FE, Kagan IA, Rimando AM (2003) Elucidation of the biosynthetic pathway of the allelochemical sorgoleone using retrobiosynthetic NMR analysis. J Biol Chem 278:28607–28611. Funa N, Ozawa H, Hirata A, Horinouchi S (2006) Phenolic lipid synthesis by type III polyketide synthases is essential for cyst formation in Azotobacter vinelandii. Proc Natl Acad Sci USA 103:6356–6361. Funabashi M, Funa N, Horinouchi S (2008) Phenolic lipids synthesized by type III polyketide synthase

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confer penicillin resistance on Streptomyces griseus. J Biol Chem 283:13983–13991. Gellerman JL, Anderson WH, Schlenk H (1976) Synthesis of anacardic acids in seeds of Ginkgo biloba. Biochim Biophys Acta 431:16–21. Miyanaga A, Funa N, Awakawa T, Horinouchi S (2008) Direct transfer of starter substrates from type I fatty acid synthase to type III polyketide synthases in phenolic lipid synthesis. Proc Natl Acad Sci USA 105:871–876. Lin LP, Sadoff HL (1968) Encystment and polymer production by Azotobacter vinelandii in the presence of beta-hydroxybutyrate. J Bacteriol 95:2336–2343. Kozubek A, Tyman JH (1999) Resorcinolic lipids, the natural non-isoprenoid phenolic amphiphiles and their biological activity. Chem Rev 99:1–26. Pan Z, Rimando AM, Baerson SR, Fishbein M, Duke SO (2007) Functional characterization of desaturases involved in the formation of the terminal double bond of an unusual 16:3D9,12,15 fatty acid isolated from Sorghum bicolor root hairs. J Biol Chem 282:4326–4335.

Reusch RN, Sadoff HL (1983) Novel lipid components of the Azotobacter vinelandii cyst membrane. Nature 302:268–270. Rock CO, Cronan JE (1996) Escherichia coli as a model for the regulation of dissociable (type II) fatty acid biosynthesis. Biochim Biophys Acta 1302:1–16. Schweizer E, Hofmann J (2004) Microbial type I fatty acid synthases (FAS): major players in a network of cellular FAS systems. Microbiol Mol Biol Rev 68:501–517. Trivedi OA, Arora P, Vats A, Ansari MZ, Tickoo R, Sridharan V, Mohanty D, Gokhale RS (2005) Dissecting the mechanism and assembly of a complex virulence mycobacterial lipid. Mol Cell 17:631–643. Wallace KK, Zhao B, McArthur HAI, Reynolds KA (1995) In vivo analysis of straight-chain and branched-chain fatty acid biosynthesis in three actinomycetes. FEMS Microbiol Lett 131:227–234. Watanabe CM, Townsend CA (2002) Initial characterization of a type I fatty acid synthase and polyketide synthase multienzyme complex NorS in the biosynthesis of aflatoxin B(1). Chem Biol 9:981–988.

33 The Biosynthesis and Evolution of Archaeal Membranes and Ether Phospholipids Y. Koga University of Occupational and Environmental Health, Munakata City, Japan [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452

2 2.1 2.2 2.3 2.4 2.5 2.6

Biosynthesis of Polar Lipids in Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 452 G-1-P Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 453 Isoprenoid-PP Formation Via Mevalonate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 Ether Bond Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 454 CDP-Activated Intermediate (CDP-Archaeol) Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . 455 Biosynthesis of Archaetidylserine and Related Phospholipids . . . . . . . . . . . . . . . . . . . . . . . 455 Saturation of the Isoprenoid Chains of Phospholipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 455

3

A Hypothetical Mechanism of Differentiation of Archaea and Bacteria by Enantiomeric Phospholipid Membrane Segregation . . . . . . . . . . . . 456

4

Conclusion and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 456

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Abstract: The biosynthetic pathway of archaeal phospholipids has been basically elucidated by means of in vitro studies. The glycerophosphate (sn-glycerol-1-phosphate, G-1-P) backbone of archaeal phospholipids, which is an enantiomer of bacterial sn-glycerol-3-phosphate (G-3-P) backbone, is formed from dihydroxyacetonephosphate by the aid of a novel enzyme, G-1-P dehydrogenase. G-1-P formation is followed by the formation of two ether bonds, activation by CDP, replacement of CMP by L-serine, and hydrogenation of the double bonds on the isoprenoid chains. The pathway resembles the bacterial phospholipid synthetic pathway in the reaction sequence. The reactions are classified into two categories. One forms the characteristic structures specific to phospholipids of either archaea or bacteria. These reactions are located in the first half of the pathway. The other one forms the common structures shared by the two domains of organisms in the last half. The enantiomeric backbone structure of glycerophosphate is the most fundamental difference between archaea and bacteria and is the features that distinguish archaea and bacteria. A hypothesis is reviewed that differentiation of archaea and bacteria had occurred based on the unique lipid structure with G-1-P and G-3-P cores.

1

Introduction

Archaeal polar lipids in general are composed of a core lipid [isoprenoid glycerol ether (archaeol or caldarchaeol, for the nomenclature of archaeal polar lipids, see Nishihara et al., 1987) and polar groups (phosphodiester-linked polar head groups or sugar moieties). The isoprenoid chains are bound at the sn-2 and 3 positions of the glycerol backbone through ether linkages and a phosphoester or a sugar group is bound at the sn-1 position (sn-glycerol1-phosphate or G-1-P structure). G-1-P is an enantiomer (a mirror image stereoisomer) of the sn-glycerol-3-phosphate (G-3-P) backbone of bacterial phospholipids. This is one of the most fundamental and definitive differences between archaeal and bacterial polar lipids, because there is no exception structurally, enzymatically or genetically in the pattern of distribution in either domain. In addition to the G-1-P backbone structure, archaeal lipids have three other unique characteristics, isoprenoid hydrocarbon chains, ether bonds, and bipolar tetraether structure. It has been interested since 1970 in the biosynthetic mechanisms by which these characteristic structures are formed. After occasional in vivo studies for 20 years, first in vitro experimental research on biosynthesis of archaeal ether lipid has been reported. Since then in this 18 years, a major route of archaeal polar lipid biosynthesis has been elucidated. A recent comprehensive review article on the biosynthesis of archaeal polar lipids (Koga and Morii, 2007) should also be referenced. The biosynthetic studies could also helped to discuss a hypothesis for differentiation of archaea and bacteria. At the end of this chapter, the hypothesis is introduced.

2

Biosynthesis of Polar Lipids in Archaea

In vitro biosynthetic studies have helped to clarify the biochemical mechanism of the formation of these unique structures of archaeal polar lipids as well as structures common to bacterial polar lipids (> Fig. 1). The pathway and the relevant enzymes and genes are briefly summarized in this section.

The Biosynthesis and Evolution of Archaeal Membranes and Ether Phospholipids

33

. Figure 1 Possible biosynthetic pathway for archaeal phospholipids. The enzymes in phospholipid biosynthesis confirmed by in vitro experiments are as follows: 1, G-1-P dehydrogenase; 2, GGGP synthase; 3, DGGGP Synthase; 4, CDP-archaeol synthase; 5, archaetidylserine synthase; 9, archaetidylinositol-P synthase; 10, archaetidylinositol-P phosphatase; 11, digeranylgeranylphospholipid reductase. Reactions 6, 7 and 8 are supported by in vivo experiments or database searches of the relevant genes. The enzymes and the reactions a and b are in the classical mevalonate pathway, while the reactions and the enzymes c and d are in the modified pathway in Methanocaldococcus jannaschii (Grochowski et al., 2006).

2.1

G-1-P Formation

How the characteristic feature of G-1-P was formed is a primary interest in the investigation of archaeal phospholipid biosynthesis. After several in vivo experiments, Nishihara and Koga (1995) found in Methanothermobacter thermautotrophicus that dihydroxyacetonephosphate (DHAP), which is supplied from the glycolysis or gluconeogenesis pathway, was directly

453

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The Biosynthesis and Evolution of Archaeal Membranes and Ether Phospholipids

reduced to G-1-P by NAD(P)H. which was catalyzed by a new enzyme, G-1-P dehydrogenase and this enzyme activity has been ubiquitously detected in the archaea so far surveyed. G-1-P dehydrogenase and the coding egsA gene are responsible for the formation of the enantiomeric glycerophosphate backbone of polar lipids specific to archaea. This enzyme is connecting gluconeogenesis (or glycolysis) with lipid biosynthesis pathway. G-3-P dehydrogenase that is present in heterotrophic archaeal cells is working for catabolism of glycerol in conjunction with glycerol kinase (G-3-P-forming) but not for lipid synthesis. Although DHAP reduction by G-1-P dehydrogenase in archaea and G-3-P dehydrogenase in bacteria with NAD(P)H for the formation of glycerophosphate is the same type of reaction, the products of the reactions are different in the stereo chemical structure. The two enzymes are fundamentally different each other in the amino acid sequences, the stereochemistry of hydride ion transfer to and from NAD(P), and the origin of the enzymes (Daiyasu et al., 2002).

2.2

Isoprenoid-PP Formation Via Mevalonate Pathway

Hydrocarbon chains of archaeal glycerol polar lipids are composed exclusively of isoprenoid derivatives in contrast to the fatty acyl chains of bacterial polar lipids. Most isoprenoid chains are made up of a C20 phytanyl chain or a C40 bisphytandiyl chain that is a head-to-head condensed dimer composed of phytanyl chains. Isoprenoid chains of archaeal phospholipids are introduced as geranylgeranyl pyrophosphate, which is formed from acetyl-CoA via the mevalonate pathway not via the DOXP pathway (Rohmer, 1999) (that was recently found to exist mainly in bacteria). Although the overall pathway is the same as or similar to the eukaryotic mevalonate pathway, a few unique aspects of the enzymes have been reported. Genes of phosphomevalonate kinase, diphosphomevalonate decarboxylase, and isopentenyl diphosphate isomerase are not orthologous to the corresponding yeast genes (Smit and Mushegian, 2000). In an archaeal species, Methanocaldococcus jannaschii, the reaction order of decarboxylation and phosphorylation of phosphomevalonate has been suggested to be inverted (Grochowski et al., 2006). Isopentenyl diphosphate isomerase was also unique in its primary structure, which was a homolog of fni gene of Streptococcus CL190 type 2 IPP isomerase.

2.3

Ether Bond Formation

G-1-P is sequentially etherified usually by geranylgeranyl pyrophosphate to form digeranylgeranyl-G-1-P (DGGGP, or unsaturated archaetidic acid). The first ether bond is formed at the sn-3 hydroxyl group of G-1-P by a soluble enzyme, 3-geranylgeranyl-sn-G-1-P synthase (GGGP synthase) (Zhang et al., 1990). This enzyme prefers G-1-P as the accepter of geranylgeranyl chain to G-3-P and was not active to DHAP or glycerol. It involves the step at which three major characteristics of archaeal polar lipid structure (G-1-P stereochemistry, ether bond, and isoprenoid chain) are assembled into one molecule. Therefore, this enzyme can be regarded as having the most important role in the establishment of archaeal lipid characteristics. GGGP synthase has been purified and the coding gene has been cloned. Furthermore, the catalytic mechanism of GGGP synthase has been reported on the basis of its crystal structure (Payandeh et al., 2006). The second ether bond is formed at the sn-2 hydroxyl group of GGGP. The product of this reaction is DGGGP or unsaturated archaetidic acid. The enzyme catalyzing this reaction is

The Biosynthesis and Evolution of Archaeal Membranes and Ether Phospholipids

33

membrane-bound, and belongs to the UbiA prenyl transferase family (Hemmi et al., 2004). The enzymes of this family except for DGGGP synthase are enzymes that transfer prenyl groups to hydrophobic ring structures such as quinones and hemes. This suggests DGGGP synthase might be derived from these prenyl transferases. DGGGP synthase is also specific to a G-1-P derived precursor.

2.4

CDP-Activated Intermediate (CDP-Archaeol) Formation

After two ether bonds are formed on the G-1-P backbone, the product, unsaturated archaetidic acid (DGGGP) is then activated by CTP to form CDP-(unsaturated) archaeol (Morii et al., 2000). Archaetidic acid and CDP-archaeol play a central role in the archaeal phospholipid biosynthetic pathway equivalent to the role of phosphatidic acid and CDP-diacylglycerol in bacterial phospholipid biosynthesis. CDP-archaeol synthase does not recognize the characteristic structure of archaeal phospholipids (the G-1-P structure or ether linkages) but is specifically active to substrates with geranylgeranyl chains. The gene for this enzyme has not been identified.

2.5

Biosynthesis of Archaetidylserine and Related Phospholipids

Archaetidylserine is synthesized from CDP-unsaturated archaeol by the replacement of CMP with L-serine (Morii and Koga, 2003). CDP-archaeol is possibly also a precursor of archaetidylmyo-inositol and archaetidylglycerophosphate, because homologs of phosphatidylserine synthase, phosphatidyl-myo-inositol synthase, and phosphatidylglycerol synthase were detected in archaea and bacteria by a BLAST search. However, among the three enzymes, only archaetidylserine synthase has been enzymatically characterized, the other two have not. These enzymes that catalyze the replacement of the CMP of CDP-archaeol by one of the various polar groups (L-serine, glycerophosphate, or myo-inositol etc.) are enzymes of CDP-alcohol phosphatidyltransferase family (Daiyasu et al., 2005). In contrast to G-1-P dehydrogenase and the ether bond-forming enzymes, CDP-alcohol phosphatidyltransferases from archaea and bacteria are very much akin to each other in terms of enzymatic activities, and those enzymes belong to the same enzyme family. This suggests that these phospholipidspolar head groups had existed before the differentiation of archaea and bacteria.

2.6

Saturation of the Isoprenoid Chains of Phospholipids

Most of the intermediates of phospholipid biosynthesis in archaea have unsaturated isoprenoids (geranylgeranyl groups), while most of the mature phospholipids have saturated isoprenoid chains. Therefore, four double bonds of a geranylgeranyl group in a phospholipid have to be hydrogenated in somewhere in the biosynthetic pathway and the responsible enzyme has been found (Nishimura and Eguchi, 2006). This enzyme can hydrogenate phospholipid intermediates with unsaturated isoprenoid chains irrespective of having a polar head group. Therefore, the exact hydrogenation step is not clear. Because NADH was required for the reduction and FAD increased the activity, NADH acts as a hydrogen donor via FAD.

455

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The Biosynthesis and Evolution of Archaeal Membranes and Ether Phospholipids

A Hypothetical Mechanism of Differentiation of Archaea and Bacteria by Enantiomeric Phospholipid Membrane Segregation

Although the phospholipid synthetic pathways in archaea and bacteria follow the same formal steps of the reaction sequence (DHAP reduction, hydrocarbon attachment, activation by CDP, and polar group attachment), it is concluded that the archaea-specific structural characteristics of phospholipids are formed in the first half of the pathway (G-1-P structure, isoprenoid chains, and ether bonds). By contrast, the common polar head groups shared by archaea and bacteria are attached in the last half of the pathway. These findings suggest that the phospholipid synthesis pathway evolved as early as the cellular stage of the common ancestor of archaea and bacteria before their differentiation. In this stage (the common ancestor is designated as being at a pre-cell stage according to Wa¨chtersha¨user, see below), membrane phospholipids had either or both kinds of the core lipids (isoprenoid-G-1-P ether, and fatty acyl G-3-P ester) and the same common polar head groups. In the very long story of the origin of life and early evolution of cells, the role of lipid membranes, especially phospholipid enantiomers, on the differentiation of archaea and bacteria was first given attention by Koga et al. (1998). Wa¨chtersha¨user (2003) has proposed a theory of the evolution of archaea and bacteria by spontaneous segregation of the isoprenoidG-1-P ether, or the fatty acyl G-3-P ester membranes in pre-cells as the common ancestor (> Fig. 2). The membrane lipids of these pre-cells must have had not only either the G-1-P or G-3-P core lipids, but also polar head groups in their membrane lipids. During the chemical evolution stages before the origin of life, the glycerophosphate that is synthesized abiotically would be expected to be racemic. The enantiomeric membranes would then be spontaneously segregated into two kinds of more homochiral membranes (G-1-P-rich and G-3-P-rich membranes). Moreover, in the course of the process of frequent collision, fusion, and fission of pre-cells, both pre-cells with more homochiral membranes and pre-cells with more heterochiral membranes arise. Wa¨chtersha¨user assumed that the more heterochiral membrane is less stable than the more homochiral membrane. Pre-cells with more heterochiral membranes would, therefore, more readily become extinct. As a result of this process, advanced pre-cells with more homochiral membranes (either G-1-P or G-3-P) appear. In addition, G-1-P dehydrogenase and G-3-P dehydrogenase evolved from different ancestral enzymes in the advanced pre-cells A (the ancestor of archaea) and the advanced pre-cells B (the ancestor of bacteria), respectively. Membrane phospholipids in the pre-cells would become completely homochiral. These are the ancestors of archaea and bacteria, respectively. Therefore, the advanced pre-cells with either homochiral core lipid carried over common polar groups on the lipid molecules and their synthesizing enzymes that had been present since the stage of pre-cells with racemic membranes. This conclusion was reached from the evolutionary consideration by Wa¨chtersha¨user (2006) and from the studies on the structure and biosynthesis of archaeal lipids by Koga and Morii (2007).

4

Conclusion and Research Needs

Although only a limited portion of the biosynthetic pathway of archaeal phospholipids has been previously elucidated, a fundamental characteristic has been brought to light.

The Biosynthesis and Evolution of Archaeal Membranes and Ether Phospholipids

33

. Figure 2 A proposed model for the differentiation of archaea and bacteria based on the spontaneous segregation of G-1-P lipid membranes and G-3-P lipid membranes. Heterochiral lipid membranes are assumed to be less stable than more homochiral membranes, and pre-cells with less stable membranes would more readily become extinct. Consequently, two kinds of pre-cells with homochiral membranes will be the ancestors of archaea and bacteria, respectively. The polar head groups that had existed at the stage of the pre-cells with racemic lipid membranes might be carried over into the both descendants, archaea and bacteria.

The pathways for archaeal and bacterial phospholipids bear a close resemblance. These are different only in synthesis of characteristic structures of archaeal and bacterial lipids. It is considered that this reflects evolutionary process of membrane phospholipids and the differentiation of archaea and bacteria. While bacterial phospholipid synthesis has been exhaustively studied, the comparatively slight research on archaeal phospholipid synthesis has nevertheless helped illuminate certain unknown features of bacterial phospholipid synthesis. For example, the unexpected activity of bacterial phosphatidylserine synthase to archaeal isoprenoid ether type serine phospholipid is shown, suggesting that the polar head groups had been shared by the membranes of the common ancestors of archaea and bacteria before differentiation. Many problems about lipid biosynthesis in archaea remain to be investigated. The in vitro biosynthesis of ethanolamine-, glycerol-, and inositol-phospholipids is urgent problems. Identification of the relevant genes and phylogenetic examination are needed for biosynthetic and evolutionary interests. Experiments that give support to the hypothesis of the differentiation of archaea and bacteria, for example, experiments using liposomes with mixed lipids of archaea and bacteria, will be highly interested. Recent results of in vitro experiments with a Methanothermobacter thermautotrophicus membrane fraction (Morii and Koga, unpublished) showed that CDP-archaeol was reacted with 1L-myo-inositol-phosphate to form archaetidyl-myo-inositol-phosphate, which was then dephosphorylated resulting in archaetidyl-myo-inositol. This pathway is a new inositolphospholipid synthetic pathway, in which the sequence of the transfer of an archaetidyl (phosphatidyl) group and dephosphorylation is inverted compared with eukaryotic phosphatidylinositol synthesis. > Fig. 1 shows this pathway.

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References Daiyasu H, Hiroike T, Koga Y, Toh H (2002) The analysis of membrane stereochemistry with the homology modeling of sn-glycerol-1-phosphate dehydrogenase. Protein Eng 15: 987–995. Daiyasu H, Kuma K, Yokoi T, Morii H, Koga Y, Toh H (2005) A study on archaeal enzymes involved in polar lipids synthesis by an approach linking the information about amino acid sequences, genomic contexts and lipid composition. Archaea 1: 399–410. Grochowski LL, Xu H, White R (2006) Methanocaldococcus jannaschii uses a modified mevalonate pathway for biosynthesis of isopentenyl diphosphate. J Bacteriol 188: 3192–3198. Hemmi H, Shibuya K, Takahashi Y, Nakayama T, Nishino T (2004) (S)-2,3-di-O-geranylgeranylglyceryl phosphate synthase from the thermoacidophilic archaeon Sulfolobus solfataricus – molecular cloning and characterization of a membrane-intrinsic prenyltransferase involved in the biosynthesis of archaeal ether-linked membrane lipids. J Biol Chem 279: 50197–50203. Koga Y, Kyuragi T, Nishihara M, Sone N (1998) Did archaeal and bacterial cells arise independently from noncellular precursors? A hypothesis stating that the advent of membrane phospholipid with enantiomeric glycerophosphate backbones caused the separation of the two lines of descent. J Mol Evol 46: 54–63. Koga Y, Morii H (2007) Biosynthesis of ether-type polar lipids in archaea and evolutionary considerations. Micobiol Mol Biol Rev 71: 97–120. Morii H, Koga Y (2003) CDP-2,3-di-O-geranylgeranylsn-glycerol: L-serine O-archaetidyltransferase (archaetidylserine synthase) in the methanogenic archaeon Methanothermobacter thermautotrophicus. J Bacteriol 185: 1181–1189. Morii H, Nishihara M, Koga Y (2000) CTP: 2,3-di-Ogeranylgeranyl-sn-glycero-1-phosphate cytidyltransferase in the methanogenic archaeon

Methanothermobacter thermoautotrophicus. J Biol Chem 275: 36568–36574. Nishihara M, Koga Y (1995) sn-Glycerol-1-phosphate dehydrogenase in Methanobacterium thermoautotrophicum: key enzyme in biosynthesis of the enantiomeric glycerophosphate backbone of ether phospholipids of archaebacteria. J Biochem 117: 933–935. Nishihara M, Morii H, Koga Y (1987) Structure determination of a quartet of novel tetraether lipids from methanobacterium thermoautotrophicum. J Biochem 101: 1007–1015. Nishimura Y, Eguchi T (2006) Biosynthesis of archaeal membrane lipids: digeranylgeranylglyceryphospholipid reductase of the thermophilic archaeon Thermoplasma acidophilum. J Biochem 139: 1073–1081. Payandeh J, Fujihashi M, Gillon W, Pai EF (2006) The crystal structure of (S)-3-O-geranylgeranylglyceryl phosphate synthase reveals an ancient fold for an ancient enzyme. J Biol Chem 281: 6070–6078. Rohmer M (1999) The discovery of a mevalonateindependent pathway for isoprenoid biosynthesis in bacteria, algae and higher plants. Nat Prod Rep 16: 565–574. Smit A, Mushegian A (2000) Biosynthesis of isoprenoids via mevalonate in archaea: the lost pathway. Genomic Res 10: 1465–1484. Wa¨chtersha¨user G (2003) From pre-cells to eukarya – a tale of two lipids. Mol Microbiol 47: 13–22. Wa¨chtersha¨user G (2006) From volcanic origins of chemoautotrophic life to bacteria, archaea and eukarya. Philos Trans R Soc Lond B 361: 1787–1808. Zhang DL, Daniels L, Poulter C (1990) Biosynthesis of archaebacterial membranes. Formation of isoprene ethers by a prenyl transfer reaction. J Am Chem Soc 112: 1264–1265.

34 Production of Wax Esters by Bacteria J.-F. Rontani Laboratoire de Microbiologie, de Ge´ochimie et d’Ecologie Marines (LMGEM-UMR 6117), Centre d’Oce´anologie de Marseille – Campus de Luminy, Marseille, France [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460

2

Different Types of Bacteria Producing Wax Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460

3

Biological Functions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 460

4 4.1 4.1.1 4.1.2 4.1.3 4.2 4.3

Formation Pathways during the Metabolism of Different Substrates . . . . . . . . . . . 461 Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 461 n-Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 461 Isoprenoid Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462 Isoprenoid Alkenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 462 n-Alkanols and Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 464 Phytol and Derivatives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 464

5

Wax Esters: A Fingerprint of the Degradation Pathways Employed by Bacteria? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 465

6

Potential Biogeochemical Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 469

7

Applications in the Manufacture of Commercial Products . . . . . . . . . . . . . . . . . . . . . . 469

8

Research Needs and Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 469

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_34, # Springer-Verlag Berlin Heidelberg, 2010

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34

Production of Wax Esters by Bacteria

Abstract: During cultivation under nitrogen-limited conditions, some prokaryotes are able to accumulate large amounts of wax esters as inclusion bodies in their cytoplasm. These lipids act as storage compounds for energy and carbon during starvation. The present review focuses on the formation pathways of these compounds during the bacterial metabolism of various substrates (hydrocarbons, alkanols, and fatty acids). A particular attention is given to the formation of isoprenoid wax esters. As a conclusion, the potential applications of these compounds are discussed.

1

Introduction

Accumulation of wax esters in intracellular lipid-bodies is a property of only a few prokaryotes. The shape of wax ester bodies is not restricted to spherical inclusions and some authors described flat, disk-like, or rectangular inclusions when the cells were cultivated on alkanes or alkanols, respectively (Wa¨ltermann and Steinbu¨chel, 2005). Some strains of Acinetobacter sp. (Makula et al., 1975) and the marine bacterium Fundibacter jadensis (Bredemeier et al., 2003) also produce extracellular wax esters, whose the function and the mechanisms of export are still unknown.

2

Different Types of Bacteria Producing Wax Esters

Wax esters are generally considered as widespread energy storage components in the genus Acinetobacter (Fixter et al., 1986). Meanwhile, accumulation of such compounds was also described for some bacterial strains belonging to the genus Moraxella, Micrococcus, Fundibacter, Neisseria, Marinobacter, and Pseudomonas but also for actinomycetes (Corynebacterium, Mycobacterium, Rhodococcus, and Nocardia) (for reviews see Ishige et al., 2003; Wa¨ltermann and Steinbu¨chel, 2005). In the particular case of Acinetobacter calcoaceticus, which was intensively studied, wax esters can reach a fraction of about 25% of the cellular dry weight (Wa¨ltermann et al., 2005).

3

Biological Functions

The main function of wax esters in bacteria is to serve as a storage compound for energy and carbon. Indeed, it was previously demonstrated that the amounts of wax esters produced increases considerably in N-limited cultures under conditions of low growth rate, where carbon and energy are in excess (Fixter et al., 1986). Thus the accumulated wax esters also appeared to be degraded to water soluble molecules and carbon dioxide during subsequent C-starvation (Fixter et al., 1986). However, other functions should also be considered: (1) lipid bodies may also act as a deposit for toxic or useless fatty acids during growth on recalcitrant carbon sources and (2) storage of evaporation-resistant lipids might also be a strategy to maintain a basic water supply in the case of desiccation (Wa¨ltermann and Steinbu¨chel, 2005).

Production of Wax Esters by Bacteria

4

Formation Pathways during the Metabolism of Different Substrates

4.1

Hydrocarbons

4.1.1

n-Alkanes

34

The formation pathways for wax ester synthesis during bacterial growth on n-alkanes were intensively studied in the case of stains belonging to the genus Acinetobacter (Ishige et al., 2000; 2003). The proposed pathways are shown in > Fig. 1. The final step of esterification is catalyzed by an acyl-CoA:alcohol transferase, which was recently characterized as a novel bifunctional wax ester synthase/acyl-CoAalcohol transferase (Kalscheuer and Steinbu¨chel, 2003). In the case of the strain Acinetobacter sp. Strain M-1 (Ishige et al., 2003), there are two possible routes for supplying alcohol: (1) by conversion of acyl-CoA to alcohol through the sequential reactions of acyl-CoA reductase and aldehyde reductase, or (2) by direct introduction from 1-alkanols or alkanals resulting from n-alkane oxidation (> Fig. 1).

. Figure 1 Proposed pathways for the production of wax esters during the metabolism of n-alkanes by Acinetobacter spp. (adapted from Ishige et al., 2003).

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Production of Wax Esters by Bacteria

. Figure 2 Proposed pathways for the production of wax esters during the metabolism of phytane by Mycobacterium ratisbonense sp. MD4. (adapted from Silva et al., 2007).

4.1.2

Isoprenoid Alkanes

Recently, the accumulation of some isoprenoid wax esters was observed during the biodegradation of phytane (2,6,10,14-tetramethylhexadecane) by the strain Mycobacterium ratisbonense sp. SD4 under N-starved conditions (Silva et al., 2007). Chemical analyses of metabolites revealed the existence of at least three different pathways for the catabolism of this isoprenoid alkane (attack of the isopropyl side, of the subterminal and terminal carbon atoms of the ethyl-terminus of the molecule) (> Fig. 2). The importance of these different degradation pathways was clearly reflected in the composition of the wax esters detected. Interestingly, triacylglycerols but not wax esters were accumulated by this strain grown on hexadecane, the unbranched homologue of phytane. The specific biosynthesis and accumulation of isoprenoid wax esters by phytane-grown cells of Mycobacterium ratisbonense sp. SD4 was attributed to the fact that branched fatty acids resulting from the catabolism of phytane may not be suitable for maintaining the functional fluidity of cellular biomembranes, and may be thus transferred to the wax ester pool.

4.1.3

Isoprenoid Alkenes

The production of 5,9,13-trimethyltetradeca-4E,8E,12-trienyl-5,9,13-trimethyltetradeca4E,8E,12-trienoate was previously observed during the aerobic degradation of squalene (2,6,10,15,19,23-hexamethyltetracosa-2,6E,10E,14E,18E,22-hexaene) by Marinobacter squalenivorans sp. nov. isolated from the marine environment (Rontani et al., 2003). A pathway

Production of Wax Esters by Bacteria

34

involving initial cleavage of the C10–C11 or C14–C15 double bonds of the squalene molecule was proposed to explain the formation of this polyunsaturated isoprenoid wax ester (> Fig. 3). This cleavage produces 5,9,13-trimethyltetradeca-4E,8E,12-trienal, which can be converted by an aldehyde dehydrogenase to the corresponding acid or reduced by an aldehyde reductase

. Figure 3 Proposed pathways for the production of wax esters during the metabolism of squalene by Marinobacter squalenivorans sp. nov. (adapted from Rontani et al., 2003).

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. Figure 4 Schematic summary of wax ester syntheses by Acinetobacteria that convert exogeneous oleic acid and other alcohol substrates into a homologous series of waxes. (adapted from Kaneshiro et al., 1996).

(alcohol dehydrogenase) to the corresponding alcohol; these two compounds being then esterified. The wax ester content increased by approximately threefold in N-limited cultures, in which the ammonium concentration corresponds to conditions often found in marine sediments. This result suggests, therefore, that the formation of isoprenoid wax esters might be favored in such environments when squalene is available.

4.2

n-Alkanols and Fatty Acids

Fixter et al. (1986) reported the occurrence of cellular inclusions with hexadecyl palmitate as principal component when Acinetobacter sp. HO1-N was cultivated on hexadecan-1-ol. The formation of wax esters was also observed during growth of the strain Acinetobacter sp. 211 on hexadecan-1-ol and olive oil (Alvarez et al., 1997). Production of wax esters by cultures of Acinetobacter strains supplemented with combined fatty (C8–C18) alcohols and acids suggested a coordinated synthesis whereby the exogenous alcohol remains unaltered, and the fatty acid is partially oxidized with removal of C2 units before esterification (Kaneshiro et al., 1996) (> Fig. 4). It was concluded that primary alcohols control chain length of the wax esters. Due to its unspecificity, the bifunctional wax ester synthase/acylCoA-diacylglycerol acyltransferase of Acinetobacter calcoaceticus ADP 1 also acts on diols (such as 1,16-hexadecanediol) affording wax diesters (Kalscheuer et al., 2003).

4.3

Phytol and Derivatives

The production of isoprenoid wax esters was also previously observed during the aerobic degradation of 6,10,14-trimethylpentadecan-2-one and phytol (3,7,11,15-tetramethylhexadec-2-en-1-ol) by Acinetobacter sp. PHY9, Pseudomonas nautica, Marinobacter sp. CAB and Marinobacter hydrocarbonoclasticus DSM 8798 isolated from the marine environment (Rontani et al., 1999a).

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In the case of 6,10,14-trimethylpentadecan-2-one, these esters result from the condensation of some acidic and alcoholic metabolites produced during the biodegradation process, which involved an initial enzymatic oxidation of the ketone to 4,8,12-trimethyltridecan-1-ol acetate (which is analogous to Baeyer-Villiger oxidation with peracids) (> Fig. 5), and seemed to operate for each of the four strains studied. The active esterase system responsible for the formation of wax esters was probably the same as that involved during hydrolysis of 4,8,12trimethyltricecan-1-ol acetate to 4,8,12-trimethyltridecan-1-ol and acetic acid (> Fig. 5). This hypothesis is well supported by the fact that 4,8,12-trimethyltridecan-1-ol constitutes the alcohol moiety of most of the wax esters identified. Although Marinobacter sp. CAB, Pseudomonas nautica and Marinobacter hydrocarbonoclasticus DSM 8798 are able to grow on 6,10,14trimethylpentadecan-2-one under denitrifying conditions, they failed to produce isoprenoid wax esters under these conditions. This was attributed to the fact that: (1) the enzymatic oxidation of ketones by way of an ester intermediate cannot operate in the absence of oxygen, and (2) the esterases involved during this process are generally inducible enzymes. The first step of the bacterial degradation of phytol involves the transient production of the corresponding aldehyde (E)-3,7,11,15-tetramethylhexadec-2-enal (phytenal). This labile compound can be converted quickly and abiotically in seawater to 6,10,14-trimethylpentadecan-2one. The production of this ketone involves addition of water to the activated double bond of phytenal followed by a retro-aldol reaction. Concurrently, the phytol can be metabolized via (E)-phytenic acid by two different pathways involving; (1) alternating b-decarboxymethylation and b-oxidation sequences or (2) hydrogenation to 3,7,11,15-tetramethylhexadecanoic acid (phytanic acid) and subsequent a- and b-oxidation. Several isoprenoid wax esters (> Fig. 6) arising from the esterification of phytol with some of its acidic metabolites were detected after growth of the different strains on this isoprenoid alcohol. Esterification activity in these cultures was not confined to 4,8,12-trimethyltridecan-1-ol; phytol appeared also to be an excellent substrate. Recently, two new isoprenoid wax ester synthases capable of synthesizing isoprenoid and acyl/isoprenoid hybrid esters were identified in Marinobacter hydrocarbonoclasticus DSM 8798 (Holtzapple and Schmidt-Dannert, 2007).

5

Wax Esters: A Fingerprint of the Degradation Pathways Employed by Bacteria?

Wax esters, which constitute useful ‘‘metabolite traps’’, appear to be particularly well suited to the study of the bacterial metabolism of isoprenoid compounds. Indeed, it is often difficult to determine the metabolic pathways employed by bacteria during the degradation of isoprenoid substrates owing to the very short lifetime of most of the metabolites formed. Wax esters, which result from the condensation of these intermediates and are accumulated by bacteria, can thus be used to monitor the metabolic pathways involved during the degradation. Analyses of isoprenoid wax esters produced by condensation of bacterial metabolites with themselves or with phytol allowed notably to show that the bacterial metabolism of phytol can be strongly modified by growth temperature and in the presence of a solid support (Rontani and Bonin, 2000) (> Fig. 7). The aerobic metabolism of phytol involves in fact the transient production of (E) phytenal, which in turn can be quickly abiotically converted to 6,10,14trimethylpentadecan-2-one or (E) phytenic acid. Low temperatures and sorption on mineral particles appeared to hinder the addition of water upon the activated double bond of (E) phytenal, which constitutes the first step of its transformation to the C18 isoprenoid ketone.

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. Figure 5 Proposed pathways for the production of wax esters during the aerobic metabolism of 6,10,14-trimethylpentadecan-2-one by marine bacteria. (adapted from Rontani et al., 1999a).

466 Production of Wax Esters by Bacteria

. Figure 6 Proposed pathways for the production of wax esters during the aerobic metabolism of phytol by marine bacteria. (adapted from Rontani et al., 1999a).

Production of Wax Esters by Bacteria

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Production of Wax Esters by Bacteria

. Figure 7 Mass fragmentograms showing the distribution of 4,8,12-trimethyltridecan-1-yl (m/z 196) and phytyl (m/z 278) wax esters (a) in free cell culture at 20 C, (b) in immobilised cell culture at 20 C and (c) in immobilized cell culture at 15 C of a marine bacterial community. (adapted from Rontani and Bonin, 2000).

Production of Wax Esters by Bacteria

34

Consequently, it was concluded that in surface temperate sediments phytol must be metabolised mainly through (E) phytenic acid and that results obtained in vitro with free cell cultures could not be systematically extended to the marine environment.

6

Potential Biogeochemical Applications

Isoprenoid wax esters are rarely abundant in sediments, probably due to their rapid hydrolysis during early diagenesis. However, it is interesting to note that despite the occurrence of these hydrolytic processes, Cranwell (1986) detected phytylphytenate in lacustrine sediments up to 40,000 years old. Similarly, esters of sterols are thought to be more stable in sediments than the corresponding free compounds, suggesting that, under some conditions, esterification can enhance the preservation potential of labile compounds such as phytol. There is a demonstrable need to identify bacterial metabolites that have sufficient structural specificity to act as biological markers for microbial degradation in the aquatic environment. Some of the isoprenoid wax esters produced during the bacterial metabolism of phytol (Rontani et al., 1999a, b), which result from the condensation of phytol metabolites between them or with phytol could play this role. Consequently, it would be useful to search for these compounds in marine sediments and particulate matter samples.

7

Applications in the Manufacture of Commercial Products

Various types of wax ester from biological sources are widely used in the manufacture of commercial products such as cosmetics, candles, printing inks, lubricants, and coating stuffs. They are produced on a scale of 3 million tons per year. Jojoba (Simmondsia californica) oil is often used as substitute for sperm whale oil to obtain unsaturated wax esters, however its use is restricted, due to its high price. The microbial production of wax esters has significant advantages over other biological processes, as the wax ester composition can be controlled by the choice of the starting materials, growth-conditions, and strains (Ishige et al., 2003). Numerous works were carried out in order to select bacterial strains belonging to Acinetobacteria or other phyla accumulating unsaturated wax esters, which resemble those of jojoba oils. Positive results were obtained, notably in the case of Acinetobacter sp. HO1-N grown on C15–C20 n-alkanes, which accumulated wax esters containing a large percentage of mono- and diunsaturated components (Dewitt et al., 1982) and presenting a close chemical similarity to those of sperm whale or jojoba oils.

8

Research Needs and Conclusions

The author hopes that this short review has provided a useful perspective on the fascinating biosynthesis mechanisms of wax esters by bacteria. Although the results obtained in the course of recent studies have considerably increased our understanding of the biosynthesis of wax esters by bacteria, there are still many gaps in our knowledge. Further studies are required

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notably to identify and characterize the enzymes implicated in these processes. Characterization of enzyme functions will be very important not only for understanding metabolic processes in microorganisms but also could yield useful enzymes for biocatalytic applications. The determination of the function and the mechanisms of export of extracellular wax esters needs further investigation. It would be also useful to search for the presence of isoprenoid wax esters in a range of sedimentary environments in order to determine if bacterial esterification can really enhance the preservation potential of labile compounds such as phytol.

References Alvarez HM, Pucci OH, Steinbu¨chel A (1997) Lipid storage compounds in marine bacteria. Appl Microbiol Biotechnol 47: 132–139. Bredemeier R, Hulsch R, Metzger JO, Berthe-Corti L (2003) Submersed culture production of extracellular wax esters by the marine bacterium Fundibacter jadensis. Mar Biotechnol 5: 579–583. Cranwell PA (1986) Esters of acyclic and polycyclic isoprenoid alcohols: Biochemical markers in lacustrine sediments. Org Geochem 10: 891–896. Dewitt S, Ervin JL, Howesorchison D, Dalietos D, Neidleman SL, Geigert J (1982) Saturated and unsaturated wax esters produced by Acinetobacter sp. HO1-N grown on C16-C20 n-alkanes. J Am Oil Chem Soc 59: 69–74. Fixter LM, Nagi MN, McCormack JG, Fewson CA (1986) Structure, distribution and function of wax esters in Acinetobacter calcoaceticus. J Gen Microbiol 132: 3147–3157. Holtzapple E, Schmidt-Dannert C (2007) Biosynthesis of isoprenoid wax ester in Marinobacter hydrocarbonoclasticus DSM 8798: Identification and characterization of isoprenoid coenzyme A synthetase and wax ester synthases. J Bacteriol 189: 3804–3812. Ishige T, Tani A, Sakai Y, Kato N (2000) Longchain aldehyde dehydrogenase that participates in n-alkane utilization and wax ester synthesis in Acinetobacter sp. Strain M-1. Appl Environ Microbiol 66: 3481–3486. Ishige T, Tani A, Sakai Y, Kato N (2003) Wax ester production in bacteria. Curr Opin Microbiol 6: 244–250. Kalscheuer R, Steinbu¨chel A (2003) A novel bifunctional wax ester synthase/acyl-CoA:diacylglycerol acyltransferase mediates wax ester and triacylglycerol biosynthesis in Acinetobacter calcoaceticus ADP1. J Biol Chem 287: 8075–8082. Kalscheuer R, Uthoff S, Luftmann H, Steinbu¨chel A (2003) In vitro and in vivo biosynthesis of wax esters by an unspecific bifunctional wax ester synthase/ acyl-CoA:diacylglycerol acyltransferase from Acinetobacter calcoaceticus ADP1. Eur J lipid Sci Technol 105: 578–584.

Kaneshiro T, Nakamura LK, Nicholson JJ, Bagby MO (1996) Oleyl oleate and homologous wax esters synthesized coordinately from oleic acid by Acinetobacter and Coryneform strains. Curr Microbiol 32: 336–342. Makula RA, Lockwood PJ, Finnerty WR (1975) Comparative analysis of lipids of Acinetobacter species grown on hexadecane. J. Bacteriol 121: 250–258. Rontani JF, Bonin P, Volkman JK (1999a) Production of wax esters during aerobic growth of marine bacteria on isoprenoid compounds. Appl Environ Microbiol 65: 221–230. Rontani JF, Bonin P, Volkman JK (1999b) Biodegradation of free phytol by bacterial communities isolated from marine sediments under aerobic and denitrifying conditions. Appl Environ Microbiol 65: 5484–5492. Rontani JF, Bonin P (2000) Aerobic bacterial metabolism of phytol in seawater: Effect of particle association on an abiotic intermediate step and its biogeochemical consequences. Org Geochem 31: 489–496. Rontani JF, Mouzdahir A, Michotey V, Caumette P, Bonin P (2003) production of a polyunsaturated isoprenoid wax ester during aerobic metabolism of squalene by Marinobacter squalenivorans sp. nov. Appl Environ Microbiol 69: 4167–4176. Silva RA, Grossi V, Alvarez HM (2007) biodegradation of phytane (2,6,10,14-tetramethylhexadecane) and accumulation of related isoprenoid wax esters by Mycobacterium ratisbonense strain SD4 under nitrogen-starved conditions. FEMS Microbiol Lett 272: 220–228. Wa¨ltermann M, Steinbu¨chel A (2005) Neutral lipid bodies in prokaryotes: Recent insights into structure, formation and relationship to eukaryotic lipid depots. J Bacteriol 187: 3607–3619. Wa¨ltermann M, Hinz A, Robenek H, Troyer D, Reichelt R, Malkus U, Galla HJ, Kalscheuer R, Sto¨veken T, von Landenberg P, Steinbu¨chel A (2005) Mechanism of lipid body formation in bacteria: How bacteria fatten up. Mol Microbiol 55: 750–763.

35 Neutral Lipids in Yeast: Synthesis, Storage and Degradation K. Athenstaedt Institute of Biochemistry, University of Technology Graz, Graz, Austria [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 472

2 Synthesis of Neutral Lipids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 472 2.1 Synthesis of Triacylglycerols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 473 2.2 Synthesis of Steryl Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 3 Lipid Particles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 3.1 Structure of Lipid Particles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 475 3.2 Budding Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 477 4

Degradation of Triacylglycerols and Steryl Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 478

5

Key Knowledge Gaps and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 478

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_35, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Neutral lipids are defined as hydrophobic molecules lacking charged groups. In yeast triacylglycerols and steryl esters comprise the major part of neutral lipids. These storage lipids accumulate when cells are provided with an excess of nutrients. Since substantial amounts of neutral lipids cannot be incorporated into biomembranes, they are sequestered from the cytosolic environment in so-called lipid particles (lipid droplets). Upon requirement storage lipids are mobilized from this compartment by triacylglycerol lipases and steryl ester hydrolases. The respective degradation products serve as energy sources and/or building blocks for membrane formation. In this chapter the reader is introduced to different mechanisms of triacylglycerol- and steryl ester synthesis, storage of these lipids in lipid particles and their subsequent mobilization. Finally, major gaps in our current knowledge about neutral lipid metabolism and research needs for a better understanding of neutral lipid turnover are highlighted.

1

Introduction

Currently, there are major efforts to gain a better understanding of the role of neutral lipids in cell metabolism, their formation and degradation, and the regulation of these processes. From the medical point of view, a detailed knowledge about neutral lipid turnover is required to combat several severe diseases of modern civilization such as obesity, arteriosclerosis and diabetes type II which are caused by detrimental accumulation of neutral lipids. Thus, medical studies are aimed at finding targets to prevent the excessive accumulation of neutral lipids and/ or at increasing the mobilization of these molecules. Since neutral lipid metabolism is well conserved from yeast to humans, many examples exist where insights in this process with the yeast system provided essential information required for a better understanding of lipid turnover in higher eukaryotes. Thus, the model organism yeast has been proven to be a highly valuable tool in biomedical research. In contrast to the medical field, the aim of biotechnological studies related to neutral lipid metabolism is to find ways to increase the amount of storage lipids, because in industry microorganisms such as the oleaginous yeast Yarrowia lipolytica are used for, e.g., single cell oil production and production of nutrients enriched in essential fatty acids. Furthermore, in times of increasing costs for fossil fuels the idea of producing biodiesel from triacylglycerols of microorganisms becomes more and more attractive.

2

Synthesis of Neutral Lipids

Triacylglycerols (TAG) and steryl esters (SE) form the main part of storage lipids in yeast. These molecules serve as ‘‘spare parts store’’ for sterols, diacylglycerols and fatty acids which are used as building blocks for membrane formation and/or for energy production. However, only recently another aspect of neutral lipid formation came into focus: synthesis of neutral lipids prevents lipotoxicity which is caused by the accumulation of excessive amounts of free fatty acids within the cell. Whereas in SE only one acyl chain is bound to the hydroxyl group at position C3 of the sterol moiety, TAG contain three fatty acids esterified to the glycerol backbone (> Fig. 1).

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. Figure 1 Structures of triacylglycerol, some of its precursors and ergosterol ester. R: Carbohydrate chain.

2.1

Synthesis of Triacylglycerols

Due to the importance of TAG as an inert storage form for fatty acids its synthesis seems to be well conserved among the different kingdoms of life. De novo synthesis of TAG includes the formation of phosphatidic acid (PtdOH), a key intermediate of glycerolipid metabolism (> Fig. 2). In yeast as well as in mammalian cells PtdOH can be synthesized by two different pathways, namely the glycerol-3-phosphate pathway and the dihydroxyacetone phosphate pathway, both named after their respective precursor (for a review see Athenstaedt and Daum, 1999). Whereas the former pathway comprises two subsequent acylation reactions yielding PtdOH, synthesis via the dihydroxyacetone phosphate pathway is a three-step process – acylation of the precursor dihydroxyacetone phosphate, reduction of the intermediate 1-acyl-dihydroxyacetone phosphate to 1-acyl-glycerol-3-phosphate (lyso-PtdOH) and a final acylation reaction yielding PtdOH. Alternatively to de novo synthesis, PtdOH can be formed from glycerophospholipids through the action of phospholipase D. Subsequent dephosphorylation of PtdOH leads to diacylglycerol, which is the direct precursor for TAG synthesis. Alternative ways for the formation of diacylglycerol include the degradation of glycerophospholipids via the action of phospholipase C and the reverse reaction of TAG synthesis, namely deacylation of TAG. However, diacylglycerol is also a substrate for the formation of

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. Figure 2 Pathways leading to the formation of triacylglycerols. De novo synthesis of triacylglycerols (TAG) includes the formation of phosphatidic acid (PtdOH). Subsequent dephosphorylation of PtdOH yields diacylglycerol (DAG), which is the direct precursor for TAG formation. TAG synthesis can be accomplished independent of acyl-CoA by transferring a fatty acid either from a phospholipid (upper arrow) or a DAG molecule (middle arrow; broken line) to DAG. The latter mechanism has not been reported in yeast. The lowest arrow shows acylation of DAG via the acyl-CoA dependent mechanism. CDP-DAG: cytidindiphosphate-diacylglycerol; Cho: choline; di-PtdGro: cardiolipin; Etn: ethanolamine; GrnP: dihydroxyacetone phosphate; Gro3P: glycerol-3phosphate; lyso-PL: lyso-phospholipid; MAG: monoacylglycerol; P-head group: phosphorylated head group of a phospholipid; PL: glycerophospholipid; PtdCho: phosphatidylcholine; PtdEtn: phosphatidylethanolamine; PtdGro: phosphatidylglycerol; PtdIno: phosphatidylinositol; PtdSer: phosphatidylserine.

phosphatidylethanolamine and phosphatidylcholine via the Kennedy pathway, and activation of PtdOH by CTP yields CDP-diacylglycerol which is the precursor for the synthesis of all glycerophospholipids (reviewed in Kent, 1995). The final step of TAG synthesis is presented by acylation of its direct precursor diacylglycerol (> Fig. 2). Mechanistically, this step occurs either by an acyl-CoA dependent or an acylCoA independent reaction. Enzymes catalyzing TAG synthesis via the former mechanism are present in all TAG synthesizing organisms. Thus, the acyl-CoA dependent acylation of diacylglycerol may be regarded as the ancestral mechanism for TAG formation. In addition, yeasts form TAG via an acyl-CoA independent reaction using the acyl-group of a glycerophospholipid as substrate. The respective enzyme catalyzes the direct transfer of a fatty acid from

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. Figure 3 Pathways leading to the formation of steryl esters. In yeast, steryl esters (SE) are formed via an acyl-CoA dependent reaction (lower arrow). A second mechanism for SE formation which is restricted to higher eukaryotes uses a phospholipid as acyl-donor (upper arrow; broken line).

the sn-2 position of a glycerophospholipid to diacylglycerol, thus combining the function of a phospholipase A2 (B) and an acyltransferase. Another acyl-CoA independent mechanism for TAG synthesis is represented by the transacylase reaction converting two diacylglycerol molecules to TAG and monoacylglycerol. However, enzymes catalyzing this type of reaction seem to have evolved during later periods, since TAG synthesis via this mechanism has only been described for higher eukaryotes, e.g., mammals and plants (Buhman et al., 2002; Lung and Weselake, 2006), but not for yeast.

2.2

Synthesis of Steryl Esters

Steryl esters are not only formed as a sterol-backup required for membrane formation when nutrients are no longer provided by the environment, but also to maintain an equilibrated sterol level within the cell. Since sterols are important membrane components which modulate the physical properties of a bilayer both excess and lack of free sterols is detrimental for the cell. In principal, similar to TAG also SE can be formed either by an acyl-CoA dependent or an acyl-CoA independent reaction (> Fig. 3). However, the latter mechanism is restricted to higher eukaryotes (mammals and plants) and has not been described for yeast. Thus, in yeast the acylation of the hydroxyl group of ergosterol and its precursors requires the presence of acyl-CoA.

3

Lipid Particles

3.1

Structure of Lipid Particles

Since substantial amounts of neutral lipids, mainly TAG and SE, cannot be incorporated into biomembranes, these lipids are sequestered in so-called lipid particles (lipid droplets). These

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storage compartments consist of a hydrophobic core which is encompassed from the cytosolic environment by a phospholipid monolayer. Few proteins embedded into the monolayer comprise the third component of lipid particles (> Fig. 4). The neutral lipid composition of the hydrophobic core of lipid particles is highly dependent on the source of this compartment. The ratio of TAG to SE differs not only between the different kingdoms of life, but also between different yeast species. Whereas nearly equal amounts of TAG and SE are forming the hydrophobic core of lipid particles of the budding yeast Saccharomyces cerevisiae (Leber et al., 1994), TAG are the main component of the respective counterparts in the oleaginous yeast Yarrowia lipolytica (Athenstaedt et al., 2006). Thus, lipid particles of the latter yeast rather resemble the respective cell compartments in adipocytes, equally storing mainly TAG, than that of the budding yeast. Structural investigations of lipid particles of Saccharomyces cerevisiae revealed that TAG and SE are not randomly distributed in the hydrophobic core of the particle but well organized (Czabany et al., 2008). Directly beneath the phospholipid monolayer there are several layers of SE further sheltering an inner core of TAG. Similar to the ratio of TAG to SE forming the hydrophobic core, also size and number of lipid particles vary and depend on the source. The average diameter of lipid particles from wild-type Saccharomyces cerevisiae cells ranges between 0.3 and 0.4 mm, although also few larger droplets with a diameter of 1.2 to 1.6 mm have been observed (Leber et al., 1994). Comparison of the lipid particle size between the oleaginous yeast Yarrowia lipolytica and the budding yeast grown under the same conditions reveals a two fold bigger size of the particles in the former yeast. Shifting Yarrowia lipolytica cells to oleic acid containing medium increases the mean diameter of the particles from 0.65 to 2.5 mm. In parallel to the size, also the neutral lipid composition and protein content changes (Athenstaedt et al., 2006) demonstrating a major influence of environmental conditions on lipid particles. Even though lipid particles of different sources vary in neutral lipid composition and size, the total amount of lipids always constitutes more than 95% of the particle weight with the remaining portion representing proteins.

. Figure 4 Model of a lipid particle. The inner center of the hydrophobic core of a lipid particle is formed of triacylglycerols (TAG) which is circumvented by several layers of steryl esters (SE) and a final phospholipid monolayer. Few proteins embedded into the monolayer comprise the third component of lipid particles.

Neutral Lipids in Yeast: Synthesis, Storage and Degradation

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35

Budding Model

During the last few years our knowledge about polypeptides involved in neutral lipid metabolism immensely increased, however, the link between neutral lipid synthesis occurring in the endoplasmic reticulum (ER) and deposition of the formed molecules in the core of a lipid particle is still missing. Currently, the most accepted model for lipid particle biogenesis is the so-called budding model (> Fig. 5). In this model, it is hypothesized that neutral lipid forming enzymes localized to the ER cluster in certain regions of this compartment and catalyze the formation of TAG and SE (> Fig. 5a). As already mentioned above, substantial amounts of these neutral lipids cannot be incorporated into the phospholipid bilayer and are therefore deposited between the two leaflets of the bilayer forming a microdroplet (> Fig. 5b, c). Ongoing synthesis and deposition of neutral lipids lead to the formation of a knob which, after reaching a ‘‘critical’’ size, buds off as a mature lipid particle (> Fig. 5d). The point of lipid particle origin may be selected by the presence of only a limited number of proteins containing either none or just a limited

. Figure 5 Budding model for lipid particle biogenesis. (a) Phospholipids (PL) or acyl-CoA serve as a fatty acid donor for diacylglycerol (DAG) acylation yielding triacylglycerols (TAG). In yeast steryl esters (SE) are exclusively formed in presence of acyl-CoA. Both neutral lipids do not fit into the phospholipid bilayer and are thus deposited between the two leaflets of the bilayer (b). Ongoing neutral lipid synthesis leads to the formation of a bud (c) which buds off of the endoplasmic reticulum (ER) as a mature lipid particle (LP) after reaching the critical size (d).

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number of transmembrane spanning regions, thus allowing formation of a neutral lipid knob (Athenstaedt et al., 1999; for recent reviews see Athenstaedt and Daum, 2006; Czabany et al., 2007; Rajakumari et al., 2008). Observations such as (1) the presence of TAG and SE synthesizing enzymes in the ER, (2) dual localization of characteristic lipid particle proteins in the particle and the ER, (3) restriction of all characteristic lipid particle proteins to the ER in a mutant which lacks neutral lipid forming enzymes and thus lipid particles, and (4) all lipid particle proteins identified so far contain none or just a limited number of transmembrane spanning domains and participate in lipid metabolic processes, support the hypothesis that lipid particles are formed by a budding process. Alternatively, neutral lipids may reach the core of a pre-existing lipid particle with the aid of a lipid transfer protein, by vesicle transport and/or membrane contact.

4

Degradation of Triacylglycerols and Steryl Esters

It would not make sense if neutral lipids once formed to store the surplus of nutrients could not be mobilized upon requirement. Thus, hydrolytic enzymes catalyzing the degradation of TAG and SE exist. The breakdown products of TAG are diacylglycerol and a free fatty acid. These two molecules serve either as building blocks for membrane lipid formation or as signaling molecules. In addition, fatty acids can be degraded by b-oxidation supplying the cell with energy. Sterols which are set free upon hydrolysis of SE can be directly or after further conversion incorporated into membranes. Furthermore, like fatty acids and diacylglycerols also sterols have the capacity to act as signaling molecules. The fatty acid simultaneously yielded by SE break down can be again channeled either into anabolic or catabolic processes. Enzymes catalyzing the hydrolysis of SE are so-called SE hydrolases and TAG lipases catalyze the degradation of TAG.

5

Key Knowledge Gaps and Research Needs

Although research of the past decade immensely contributed to our current understanding of neutral lipid turnover in yeast and in the following in higher eukaryotes, many aspects of neutral lipid synthesis, storage and degradation require further investigation. Studies with higher eukaryotic organisms (mammals, plants) demonstrated that TAG can be formed from two diacylglycerol molecules by a transacylation reaction (Buhman et al., 2002; Lung and Weselake, 2006). For the budding yeast Saccharomyces cerevisiae and the fission yeast Schizzosaccharomyces pombe such a mechanism for TAG synthesis has not been described. Both yeast species accumulate rather moderate amounts of TAG (neutral lipids) compared to oleaginous yeasts. A mutant of the oleaginous yeast Yarrowia lipolytica deleted of genes homologous to major TAG synthases of the budding yeast (fission yeast) lacks TAG synthesis via both the acyl-CoA dependent pathway and the phospholipid:diacylglycerol acyltransferase pathway but still contains TAG (K. Athenstaedt, unpublished result). One possible explanation is that this yeast harbors (an) enzyme(s) catalyzing the transfer of fatty acids between two molecules of diacylglycerol thus yielding TAG. The concerted action of TAG synthases catalyzing the formation of this neutral lipid via all three mechanisms (see > Fig. 2) may account for the higher accumulation of TAG in Yarrowia lipolytica compared to

Neutral Lipids in Yeast: Synthesis, Storage and Degradation

35

non-oleaginous yeast species. The factors which are indeed causing the excessive accumulation of TAG in oleaginous yeasts remain to be elucidated. To date, the most accepted model for lipid particle formation is the budding model described above. Results of several investigations concerning lipid particles are pinpointing to a relationship between this cell compartment and the ER. However, despite major efforts to unravel the enigma of lipid particle origin, direct evidence for a ‘‘budding off of the ER’’ mechanism is still missing. In higher eukaryotes neutral lipids are not only stored within the cell as lipid particles, but also secreted in form of lipoproteins. The budding model for lipid particle biogenesis describes a budding process to the cytosolic side of the ER membrane. In contrast, lipoproteins are thought to bud to the luminal side of the phospholipid bilayer of the ER. If lipid particles are indeed formed by a budding process one question which has to be posed is which factors determine the sidedness of the budding process? Are helper proteins required for the budding off of the ER membrane? Given that such an auxiliary protein facilitating the budding process indeed exists, it remains most probably in the ER because all single mutants deleted of a lipid particle protein contain lipid particles (Fei et al., 2008). Another aspect of lipid particle formation which remains to be elucidated is why lipid particles of different sizes are observed within one cell. One possible explanation is that the size of the particle is determined by the moment of budding from the ER. Alternatively, lipid particles of similar size may bud off the ER and subsequently increase their diameters by incorporating additional neutral lipids, thus leading to small ‘‘young’’ and huge ‘‘old’’ particles. The incorporation of additional material into the hydrophobic core of lipid particles may occur by, e.g., vesicle transport, membrane contact and/or de novo synthesis of lipids on the particle. Evidence for the occurrence of such mechanisms already exist and require further investigation. As an example, by shifting cells of the oleaginous yeast Yarrowia lipolytica from glucose to oleic acid containing medium, not only the size but also the neutral lipid composition of lipid particles is altered (Athenstaedt et al., 2006). In addition, this shift complements the ‘‘standard’’ lipid particle proteome with some proteins involved in targeting and fusion processes of transport vesicles. The hypothesis that the size of a particle increases through de novo synthesis of neutral lipids catalyzed by lipid particle resident enzymes is supported by the fact that the budding yeast contains the whole set of enzymes required for de novo synthesis of TAG except a phosphatidate phosphatase. However, a phosphatidate phosphatase is present in the cytosol and has therefore access to its substrate PtdOH formed by enzymes localized to the lipid particle. Furthermore, the set of TAG synthesizing enzymes may also provide a means to compensate random TAG degradation which may occur, since the lipid particle proteome contains also several neutral lipid degrading enzymes (See also > Chapter 41 Players in the neutral lipid game – proteins involved in neutral lipid metabolism in yeast, this volume). In brief, even though at first sight our knowledge about different mechanisms contributing to the formation and maintenance of lipid particles in yeast seems to be complete, some aspects remain to be elucidated. One of the biggest challenges in the future will be unraveling the enigma of lipid particle biogenesis. Especially answering the question concerning the origin of lipid particles will be a major step forward for our understanding of neutral lipid metabolism not only in yeast but also in higher eukaryotes. For further information about neutral lipid metabolism the interested reader is referred to (See also > Chapter 41 Players in the neutral lipid game – proteins involved in neutral lipid metabolism in yeast, this volume) describing polypeptides involved in neutral lipid metabolism in yeast and some recent reviews covering specifically the topic of neutral lipid metabolism in the model organism yeast (Czabany et al., 2007; Rajakumari et al., 2008).

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References Athenstaedt K, Daum G (1999) Phosphatidic acid, a key intermediate in lipid metabolism. Eur J Biochem 266: 1–16. Athenstaedt K, Daum G (2006) The life cycle of neutral lipids: synthesis, storage and degradation. Cell Mol Life Sci 63: 1355–1369. Athenstaedt K, Jolivet P, Boulard C, Zivy M, Negroni L, Nicaud J-M, Chardot T (2006) Lipid particle composition of the yeast Yarrowia lipolytica depends on the carbon source. Proteomics 6: 1450–1459. Athenstaedt K, Zweytick D, Jandrositz A, Kohlwein SD, Daum G (1999) Identification and characterization of major lipid particle proteins of the yeast Saccharomyces cerevisiae. J Bacteriol 181: 6441–6448. Buhman KK, Smith SJ, Stone SJ, Repa JJ, Wong JS, Knapp FF Jr., Burri BJ, Hamilton RL, Abumrad NA, Farese RV Jr. (2002) DGAT1 is not essential for intestinal triacylglycerol absorption or chylomicron synthesis. J Biol Chem 277: 25474–25479. Czabany T, Athenstaedt K, Daum G (2007) Synthesis, storage and degradation of neutral lipids in yeast. Biochim Biophys Acta 1771: 299–309.

Czabany T, Wagner A, Zweytick D, Lohner K, Leitner E, Ingolic E, Daum G (2008) Structural and biochemical properties of lipid particles from the yeast Saccharomyces cerevisiae. J Biol Chem 283: 17065–17074. Fei W, Alfaro G, Muthusamy B-P, Klaasen Z, Graham TR, Yang H, Beh CT (2008) Genome-wide analysis of sterol-lipid storage and trafficking in Saccharomyces cerevisiae. Eukaryt Cell 7: 401–414. Kent C (1995) Eukaryotic phospholipid biosynthesis. Annu Rev Biochem 64: 315–343. Leber R, Zinser E, Zellnig G, Paltauf F, Daum G (1994) Characterization of lipid particles of the yeast, Saccharomyces cerevisiae. Yeast 10: 1421–1428. Lung SC, Weselake RJ (2006) Diacylglycerol acyltransferase: a key mediator of plant triacylglycerol synthesis. Lipids 41: 1073–1088. Rajakumari S, Grillitsch K, Daum G (2008) Synthesis and turnover of non-polar lipids in yeast. Prog Lipid Res 47: 157–171.

Part 6

Genetics of Biogenesis

36 Methanogenesis M. Rother Institut fu¨r Molekulare Biowissenschaften, Molekulare Mikrobiologie & Bioenergetik, Johann Wolfgang Goethe-Universita¨t Frankfurt am Main, Frankfurt, Germany [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 484 2 Hydrogenotrophic Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 486 3 Methylotrophic Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 492 4 Acetotrophic Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 494 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 495

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_36, # Springer-Verlag Berlin Heidelberg, 2010

484

36

Methanogenesis

Abstract: The biological formation of methane, methanogenesis, is an essential link in the global carbon cycle. Organic matter is decomposed anaerobically into carbon dioxide (plus hydrogen), formate, and acetate, which would, if not reduced to gaseous methane, otherwise accumulate. The only organisms producing significant amounts of methane are methanogenic archaea, strictly anaerobic members of the Euryarchaeota. All methanogens investigated thus far depend on the process of methanogenesis for energy conservation and most of them are only capable to reduce CO2 with H2 or formate to methane. In these so called hydrogenotrophic methanogens, all factors involved are therefore essential and regulation of the respective genes is moderate, mostly affecting differential expression of isogenes and fine tuning of gene expression in response to growth phase and/or substrate concentration. However, many members of the order Methanosarcinales have a broader substrate spectrum and are able to grow with, beside H2 + CO2, methylated compounds, such as methanol, methylamines, or methyl-sulfides, and acetate. Methanogenesis from these substrates employs distinct yet overlapping pathways. Consequently, expression of the genes encoding one of the pathways is very stringently regulated in response to substrate availability, its concentration, and other substrates present. This section aims at summarizing current knowledge about the organization and regulation of expression of genes encoding factors involved in the different pathways of methanogenesis.

1

Introduction

Methane is the most abundant hydrocarbon present in our atmosphere. Its current concentration of ca. 1750 ppb (Dlugokencky et al., 2003, and see http://www.epa.gov/methane) has more than tripled since pre-industrial times and an estimated 5
1014 g of biologically produced methane are annually released into our atmosphere. Methane represents either a highly attractive renewable energy source because aerobic combustion of methane produces only H2O and CO2, or a frightening contribution to global warming because the global warming potential of methane is more than 20 times higher than that of CO2 (http://www. ipcc.ch). The only organisms producing (significant amounts of) methane are methanogenic archaea. In anaerobic environments lacking abundant electron acceptors such as sulfate or nitrate the final breakdown product of organic matter is methane, connecting anaerobic with aerobic environments. Thus, methanogenesis, the biological formation of methane, plays an essential role in the global carbon cycle (Ferry and Lessner, 2008). All methanogenic archaea investigated to date strictly rely on methanogenesis for energy conservation and, thus, growth. Principally, methanogenic substrates are converted to methane with the concomitant generation of an ion motive force across the cytoplasmic membrane, which can then be employed for ATP synthesis or other energy requiring cellular processes. The number of substrates utilized for methanogenesis is quite limited reflecting the narrow ecological niche methanogens occupy: most methanogens are only able to grow with H2 + CO2 (or formate; hydrogenotrophs, > Fig. 1), some can utilize methylated compounds like methanol and methylamines (methylotrophs, > Fig. 2), and some can grow with acetate (acetotrophs, J. G. Ferry, > Chapter 23, Vol. 1, Part 5, > Fig. 3). These different substrate classes are metabolized via three distinct, but overlapping, pathways of methanogenesis (for reviews, see Ferry, 1993; Keltjens and Vogels, 1993; Thauer, 1998; Deppenmeier et al., 1999). While most members of the Methanobacteriales and Methanococcales are obligate hydrogenotrophs, many members of the Methanosarcinales can also grow either methylotrophically or

Methanogenesis

36

. Figure 1 Model for the path of hydrogenotrophic methanogenesis in Methanosarcina. CM, cytoplasmic membrane; CoM, coenzyme M; CoM-S-S-CoB, heterodisulfide; ECH, energy converting hydrogenase; F420, cofactor F420; F420H2, reduced F420; Fd, ferredoxin; FdH2, reduced ferredoxin; FMD, formyl-methanofuran dehydrogenase; Frh, F420-dependent hydrogenase; FTR, formyl-MF: H4SPT formyltransferase; H4SPT, tetrahydrosarcinapterin; HDR, heterodisulfide reductase; MCH, N5,N10-methenyl-H4SPT cyclohydrolase; MCR, methyl-CoM reductase; MER, N5, N10-methylene-H4SPT reductase; MF, methanofuran; MTR, N5-methyl-H4SPT:CoM methyltransferase; P, methanophenazine; Vho, F420-nonreducing hydrogenase; see text for details; modified from (Deppenmeier et al., 1996; Deppenmeier and Mu¨ller 2007).

acetotrophically, or both (See > Chapter 42–47, Vol. 1, Part 7). In the former two groups all factors involved in methanogenesis are essential and regulation of the respective genes is moderate, mostly affecting differential expression of isogenes and fine-tuning of gene expression in response to growth phase and/or substrate concentration. The latter group, on the other hand, needs to stringently regulate methanogenesis-related genes in response to substrate availability, its concentration, and other substrates present, in order to maximize the amount of energy that can be conserved (acetate, for example, allows only for approximately 1/4 of the energy to be conserved as compared to methanol) and to avoid unnecessary protein

485

486

36

Methanogenesis

. Figure 2 Model for the path of methylotrophic methanogenesis from methanol in Methanosarcina. ?, unknown path of FdH2 oxidation; [Co], corrinoid protein of MT1; FPO, F420H2 dehydrogenase; MT1, methanol:cob(I)alamin methyltransferase; MT2, methyl-corrinoid:CoM methyltransferase; see legend to > Fig. 1 for other abbreviations and text for details.

synthesis. This chapter reviews current knowledge about organization - and regulation of expression - of genes encoding factors, which are employed in the different pathways of methanogenesis. > Table 1 summarizes the key-findings of three transcription analyses in regard to methanogenesis-related genes (Morgan et al., 1997; Hovey et al., 2005; Hendrickson et al., 2007).

2

Hydrogenotrophic Methanogenesis

Most methanogens known possess the hydrogenotrophic pathway, in which CO2 is sequentially reduced to methane in seven steps via coenzyme-bound intermediates using H2 as the electron donor (> Fig. 1). CO2 is first reduced to the formyl-level and attached to methanofuran (MF, a 2-aminomethylfuran derivative) by formyl-MF dehydrogenase (FMD, Bobik et al., 1990). The electron donor for FMD was shown to be reduced ferredoxin in

Methanogenesis

36

. Figure 3 Model for the path of acetotrophic (aceticlastic) methanogenesis in Methanosarcina. The role of factors in grey is either uncertain or unknown; Ack, acetate kinase; [CO], CODH/ACS-bound CO; CODH/ACS, carbon monoxide dehydrogenase/acetyl-CoA synthase; HS-CoA, coenzyme A; Pta, phosphotransacetylase; ?, unknown oxidoreductase coupling FdH2 (or H2?) oxidation to methanophenazine reduction; see legend to > Fig. 1 for other abbreviations and text for details.

Methanosarcina (Meuer et al., 2002). A membrane-bound, energy converting hydrogenase (ECH, Kunkel et al., 1998), homologous to hydrogenase 3 from Escherichia coli, couples the oxidation of H2 to the reduction of ferredoxin. This endergonic reaction is driven by the ion motive force (Hedderich and Forzi, 2005). Genomic analysis suggests that in obligate hydrogenotrophs formyl-MF is generated by a similar mechanism (Hendrickson et al., 2004). In Methanosarcina barkeri and Methanosarcina mazei ECH is encoded by the echABCDEF operon but its expression-level under hydrogenotrophic conditions, compared to other substrates, has so far not been reported. Methanococcus maripaludis and Methanothermobacter marburgensis, obligate hydrogenotrophs, encode two isoforms of ECH, designated EHA and EHB, which contain at least 16 subunits (Tersteegen and Hedderich, 1999). Mutational and transcription analysis suggests that EHA functions in H2-dependent ferredoxin reduction

487

F420-dependent hydrogenase

 ++  ++

fruADGB

frcADGB

frhADGB



n.p.

n.p.

n.p.

n.p.





ehbEEFGJKLMO



mcrBDCGA

MCR



Conditiona Methanold

+++

ehaA-T



mtrEDCBAFGH

MTR





mer

MER

n.p.

+

n.d.



hmd

HMD

+

echABCDEF

+++

mtd

MTD

ECH



mch

MCH

+

+

+

n.p.



ftr

c

mrtBDGA



fmdECB



H2-limitation

fwdHFGDACB

Gene(s)b

FTR

FMD

Proteinb



n.p.

n.p.

n.p.

n.p.

++

n.p.

+





n.p.











Acetatee

n.d.

+++(–Se)

n.d.

n.d.

n.d.

n.d.

Growth phase, temperature, pH

n.d.

n.d.

++(–Ni)

+(–Ni)

n.d.

n.d.

++(+Mo)

n.d.

Other

Noll et al. (1999)

Bonacker et al. (1992) Pennings et al. (1997) Pihl et al. (1994)

Rother et al. (2005)

Afting et al. ( 2000)

Hochheimer et al. (1996)

Referencef

36

. Table 1 Relative mRNA abundances of genes and operons encoding proteins involved in methanogenesis

488 Methanogenesis

 n.d. n.d. n.d.  

hdrABC

hdrDE

fpoABCDHIJKLMNO

pta-ack

ascA

cdhABCDE

FPO

Pta/Ack

AMP-ACS

CODH/ACS

n.p.







+++



n.d.





n.p.

n.p.

n.p.

+++

+

+++





n.d.





n.d.

n.p. +++(–Se)

n.d.

n.d.

n.d.

n.d.

n.d.

n.d.

n.d.

n.d.

Noll et al. (1999)

Rother et al. (2005) Li et al. (2006)

Li et al. (2006)

Deppenmeier (1995)

b

Conditions leading to differential gene expression/protein synthesis; see text for details For simplicity, the methylotrophic, substrate-specific methyltransferases have been omitted c Versus leucine/phosphate limitation d Versus acetate e Versus methanol f References other than Morgan et al. (1997); Hovey et al. (2005); and Hendrickson et al. (2007)  constitutive, max. twofold increase/decrease; + 2–4-fold increase; ++ 5–10-fold increase; +++ >tenfold increase; – decrease; n.d. not determined; n.p. not present in methylo-/ acetotrophs

a

n.d.

vhtGACD

HDR

 n.d.

vhcDGAB

 

mvhDGAB

vhuDGAUB

vhoGAC

F420-nonreducing hydrogenase

Methanogenesis

36 489

490

36

Methanogenesis

during methanogenesis while EHB is involved in reducing ferredoxin for anabolic purposes (Porat et al., 2006; Hendrickson et al., 2007). The expression of both operons in both organisms is only mildly affected (app. twofold change in transcript abundance) by the availability of H2 or by the growth rate (Tersteegen and Hedderich, 1999; Hendrickson et al., 2007). The same is true for the two isoforms of FMD (Hendrickson et al., 2007). However, expression of the FMD encoding operons is influenced by the supply with tungsten and/or molybdenum because one FMD contains W and the other Mo (encoded by the fwdHFGDACB and fmdECB operons, respectively, in M. marburgensis (Schmitz et al., 1992; Hochheimer et al., 1996). The two next steps in hydrogenotrophic methanogenesis are the transfer of the formylgroup from MF to tetrahydromethanopterin (H4MPT, a cofactor functionally analogous to tetrahydrofolate; Methanosarcina species contain a modified H4MPT designated tetrahydrosarcinapterin, Van Beelen et al., 1984), catalyzed by formyl-MF:H4MPT formyltransferase (FTR), and the subsequent conversion of N5-formyl-H4MPT to N5,N10-methenyl-H4MPT catalyzed by N5,N10-methenyl-H4MPT cyclohydrolase (MCH). Both enzymes consist of single polypeptides and the encoding genes (ftr and mch, respectively) appear to be regulated at most twofold in response to the H2 partial pressure or the growth rate (Morgan et al., 1997; Hendrickson et al., 2007). All hydrogenotrophic methanogens analyzed to date contain a coenzyme F420dependent N5,N10-methylene-H4MPT dehydrogenase (MTD), which catalyzes the reduction of N5,N10-methenyl-H4MPT with reduced F420 (F420H2, a 5-deaza-ribolflavin derivative) to N5,N10-methylene-H4MPT (Schwo¨rer and Thauer, 1991). F420 is reduced by the F420-dependent hydrogenase, an oligomeric NiFe-enzyme loosely attached to the inner aspect of the cytoplasmic membrane (Thauer et al., 1993). The encoding genes are designated fruADGB and frcADGB (in Methanococcus species, see below) or frhADGB (in Methanothermobacter and Methanosarcina species). The subsequent step in hydrogenotrophic methanogenesis, reduction of N5,N10-methylene-H4MPT to N5-methyl-H4MPT, also depends on F420H2 as the electron donor and is catalyzed by N5,N10-methylene-H4MPT reductase (MER). Both the encoding mer gene, the mtd gene, and the fru (frh) genes appear to be regulated by a common mechanism in Methanothermobacter thermautotrophicus and M. maripaludis: when H2 becomes limiting expression of the three genes/operons simultaneously increases 3.5–22-fold (Morgan et al., 1997; Hendrickson et al., 2007). The reduction of N5,N10methenyl-H4MPT to N5,N10-methylene-H4MPT involving the F420-dependent hydrogenase and MTD is also catalyzed in one step by the H2-dependent N5,N10-methylene-H4MPT dehydrogenase (HMD) (Shima and Thauer, 2007), which is present in most methanogens, except those belonging to the orders Methanomicrobiales and Methanosarcinales (Thauer et al., 1993). The gene (hmd) is expressed at an elevated level under conditions of ample H2 supply and encodes a nickel- and iron-sulfur cluster-free hydrogenase (Zirngibl et al., 1992). Instead, it contains a unique Fe-coordinating pyridone derivative as cofactor (Shima et al., 2004). HMD has a lower affinity for H2 but a higher specific activity than the F420-dependent hydrogenase explaining their differential regulation. Furthermore, HMD and MTD are upregulated when nickel is limiting growth (Afting et al., 2000). It has been proposed that under these conditions MTD actually operates in the opposite direction, generating F420H2 needed for the MER-catalyzed reaction and for anabolic purposes, because the activity of F420dependent hydrogenase, a Ni/Fe-enzyme is compromised (Afting et al., 1998). The next step in hydrogenotrophic methanogenesis is the transfer of the methyl-group from H4SPT to coenzyme M (HS-CoM, 2-mercaptoethanesulfonic acid) catalyzed be the

Methanogenesis

36

membrane-integral N5-methyl-H4MPT:coenzyme M methyltransferase (MTR, Ga¨rtner et al., 1993). The function of only two of eight of its subunits, encoded by the mtrEDCBAFGH operon, is known: MtrA harbors a corrinoid prosthetic group which is methylated and demethylated in the catalytic cycle (Harms and Thauer, 1996a), MtrH was shown to exhibit methyl-H4MPT:cob(I)alamin methyltransferase activity (Hippler and Thauer, 1999). It was clearly established that MTR couples the exergonic methyl-transfer from H4MPT to HS-CoM to sodium ion extrusion and, thus, functions as a primary sodium ion pump (Becher et al., 1992), which explains why all methanogens investigated so far strictly depend on the presence of sodium ions for methanogenesis (Mu¨ller et al., 1993; Deppenmeier et al., 1996). Consequently, the mtrEDCBAFGH operon is constitutively expressed under hydrogenotrophic, methylotrophic, and acetotrophic conditions in all methanogens investigated. The ultimate step in all methanogenic pathways is the reduction of methyl-CoM to methane catalyzed by methyl-CoM reductase (MCR, Ermler et al., 1997; Thauer, 1998). Methane and heterodisulfide (CoM-S-S-CoB) are generated from methyl-CoM and coenzyme B (N-7-mercaptoheptanoyl-O-phospho-L-threonine, HS-CoB), which is the electron donor for this reaction. MCR contains cofactor F430 (a Ni-porphinoid) in the active site (Thauer, 1998) and the three subunits constituting the active enzyme are encoded together with two open reading frames by the mcrBDCGA operon. The function of McrC and McrD is unknown but it is possible that they are involved in the extensive post-translational modification of the native enzyme (Kahnt et al., 2007). Expression of the mcr genes is mostly constitutive, as expected for an essential factor. However, transcription of the operon is threefold higher in M. acetivorans cells grown on acetate as compared to methanol (Rother et al., 2005; Li et al., 2006). It was argued that elevated levels of MCR are required for sufficient turnover rates to support growth on this poor substrate. Conversely, the same conditions do not result in differential mcr expression in M. mazei (Hovey et al., 2005). Some members of the Methanobacteriales (and Methanococcus jannaschii) encode two isoforms, MCRI (mcrBDCGA) and MCRII (mrtBDGA) (Steigerwald et al., 1993, see also tigr.org), the genes of which are differentially regulated depending on growth conditions such as the temperature, pH, substrate concentration, reducing agents present, and growth phase (Bonacker et al., 1992; Pihl et al., 1994; Morgan et al., 1997; Pennings et al., 1997). CoM-S-S-CoB represents the terminal electron acceptor of an energy-conserving electron transport chain and reduction of CoM-S-S-CoB with H2 as the electron donor involves (at least) two enzymes, F420-nonreducing hydrogenase and heterodisulfide reductase (HDR) (Deppenmeier, 2002; Hedderich et al., 2005). Apparently, differences exist between the mechanisms employed by obligate (e.g., Methanothermobacter) and facultative (e.g., Methanosarcina) hydrogenotrophs (i.e., methylotrophs). Methanosarcina species employ a membrane-bound, two-subunit, cytochrome b-containing HdrDE (HdrE being the cytochrome b), which most probably accepts the electrons to form HS-CoM and HS-CoB from methanophenazine, a membrane-integral electron carrier functionally analogous to quinones (Abken et al., 1998). Expression of hdrDE in M. mazei and HdrDE protein levels in M. acetivorans vary at most twofold for all growth conditions tested (Hovey et al., 2005; Li et al., 2006). The reduction of methanophenazine by H2 is catalyzed by F420-nonreducing hydrogenase (Ide et al., 1999), which is encoded by homologous vhoGAC and vhtGACD operons, respectively (Deppenmeier et al., 1995). While the vho genes are constitutively expressed under all conditions tested, transcripts of the vht genes could not be detected in acetate-grown M. mazei (Deppenmeier, 1995). The function of vhtD is unknown; the other gene products constitute the enzyme, which is composed of a small (VhoG/VhtG)

491

492

36

Methanogenesis

and a large (VhoA/VhtA) subunit, typical for NiFe-hydrogenases, and a membrane-spanning cytochrome b (VhoC/VhtC). With the active site facing the periplasm, transfer of the electrons via methanophenazine and the two cytochrome b subunits (VhoC and HdrE) to the site of CoM-S-S-CoB reduction (HdrD), the overall reaction would lead to the generation of two scalar protons, and thus, a proton motive force (Deppenmeier, 2004). Obligate hydrogenotrophs like Methanococcus and Methanothermobacter species lack both cytochromes and methanophenazine. They reduce CoM-S-S-CoB by a cytoplasmic multienzyme complex composed of the F420-nonreducing hydrogenase (Mvh in Methanothermobacter, Vhu and Vhc in Methanococcus) and the heterodisulfide reductase HdrABC (encoded by the hdrABC operon), an iron-sulfur flavoprotein. There is currently no information about the role of the flavin in HdrABC and no evidence is available that any of the subunits could be located on the periplasmic aspect of the membrane; it is therefore completely unclear how the exergonic reduction of CoM-S-S-CoB can be coupled to energy conservation in these organisms (Deppenmeier and Mu¨ller, 2007). Mvh, Vhu and Vhc are encoded by the mvhDGAB, vhuDGAUB and vhcDGAB operons, respectively. Expression of mvhDGAB appears to be regulated be the H2 supply and follows the same pattern as the hmd and mrt genes (see above, Morgan et al., 1997). The vhu and vhc genes of M. maripaludis are only marginally regulated by H2 or the growth rate (Hendrickson et al., 2007). However, the supply of selenium has a profound impact on their expression and of that of the homologous genes in M. voltae (see below). The role of selenium in obligate hydrogenotrophs. Within the Archaea, selenoproteins, i.e., proteins containing the unusual amino acid selenocysteine (Bo¨ck et al., 2006), appear to be restricted to Methanococcus and the related Methanopyrus branch (Rother et al., 2001b). Genome sequence analyses, radioactive in vivo labeling, and mutational studies identified at least six methanogenesis-related selenoproteins in M. jannaschii, M. voltae, M. maripaludis, and Methanopyrus kandleri: Formate dehydrogenase (FdhA), F420-reducing (FruA) and F420-nonreducing (VhuD and VhuU) hydrogenases, FMD (FwuB) and HDR (HdrA) (Bult et al., 1996; Sorgenfrei et al., 1997; Vorholt et al., 1997; Wilting et al., 1997; Rother et al., 2001a; Hendrickson et al., 2004). These organisms also encode, to various degrees, isoforms where the selenocysteine residue is replaced by cysteine (Rother et al., 2001b). In M. maripaludis, for example, all of its selenoproteins are non-essential during growth with H2 + CO2 because they can be complemented by the corresponding cysteine-isoforms. Growth on formate, however, is impaired when the capacity to synthesize selenoproteins is abolished, which was attributed to the strict selenium-dependence of formate dehydrogenase in this organism (Rother et al., 2003). In M. voltae, regulation of the frc and vhc (cysteine encoding) genes (Sorgenfrei et al., 1997) was shown to involve both selenium-dependent repression and activation of transcription (Noll et al., 1999; Mu¨ller and Klein, 2001). Repression is mediated by HrsM, a LysR-type regulator (Sun and Klein, 2004). This report is, to the author’s knowledge, the only example where detailed information about the mechanism of regulation of any gene involved in hydrogenotrophic methanogenesis is available. However, how the selenium-status is sensed in M. voltae and how this signal is transduced, is not known.

3

Methylotrophic Methanogenesis

Most methanogenic archaea capable of growing with methylated substrates as the sole source of energy are members of the Methanosarcinales with M. barkeri, and M. mazei, being the best-studied methylotrophic methanogens. Recently, M. acetivorans has also become an

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36

important model for studying methanogenesis because it is the first Methanosarcina for which a genetic system has been established (Metcalf, 1999; Rother and Metcalf, 2005). Many Methanosarcina species can use numerous methylated compounds for growth, including methanol, trimethylamine (TMA), dimethylamine (DMA), monomethylamine (MMA), and dimethylsulfide (Hippe et al., 1979; Mah, 1980; Van der Maarel and Hansen, 1997). The methylotrophic pathway starts by funneling the substrates into the methanogenic pathway via activation by two substrate-specific methyltransferases designated MT1 and MT2 (Ferry, 1999, > Fig. 2). The substrate-specific MT1 consists of two components present in a 1:1 ratio: a corrinoid protein (MtaC for methanol, MtmC for MMA, MtbC for DMA, MttC for TMA) and a methyltransferase (MtaB for methanol, MtmB for MMA, MtbB for DMA, MttB for TMA) that transfers the methyl group from the substrate to the prosthetic group of the corrinoid protein. Subsequently, the second methyltransferase MT2 (MtaA for methanol, MtbA for methylamines) transfers the methyl group from the corrinoid protein to HS-CoM. MethylCoM is then disproportionated in a 3:1 ratio; one mole of methyl-CoM is oxidized to CO2 for every three moles of methyl-CoM reduced to methane. Methyl-CoM oxidation occurs through a reverse of the CO2 reduction pathway, generating reducing equivalents for methyl reduction (Keltjens and Vogels, 1993). However, how reduced ferredoxin accruing in the last reaction is re-oxidized (see > Fig. 2) is somewhat uncertain because a M. barkeri mutant lacking ECH grows indistinguishable from the wild type with methanol and generates methane and CO2 with wild-type rates (Meuer et al., 2002). The genes for MT1 usually comprise an operon, while MT2 is transcribed monocistronically (Harms and Thauer, 1996b; Sauer et al., 1997). Interestingly, the genes encoding MT1 forms that are required for the use of methanol, TMA, DMA and MMA, respectively, are present in two or three copies each, whereas the corresponding MT2 is encoded only in single copy. This pattern of ‘‘gene redundancy’’ is conserved across each of the sequenced Methanosarcina genomes (Deppenmeier et al., 2002; Galagan et al., 2002; Maeder et al., 2006). Notably, all methylamine-dependent MT1 isoforms were shown to depend during translation of their respective mRNAs on the suppression of in-frame stop-codons with the unusual amino acid pyrrolysine (Krzycki, 2005; Mahapatra et al., 2006). Evidence that each isoform of a particular substrate-specific MT1 serves a specific function was obtained through mutational studies (Pritchett and Metcalf, 2005) and transcription analyses, employing promoter/reporter gene fusions and DNA-microarrays (Hovey et al., 2005; Veit et al., 2005; Bose et al., 2006). Transcriptome analysis of M. mazei showed that of the methanol-specific MT1-encoding genes (mtaCB1, mtaCB2, and mtaCB3) mtaCB1 was expressed on methanol at a 10–30-fold higher level than on acetate. Expression of mtaCB2, on the other hand, was only observed in acetate-grown cells. Transcription of mtaCB3 could not be detected under any condition (Hovey et al., 2005). In contrast, a study employing fusions of the three mtaC promoters with a reporter gene in M. acetivorans revealed a more complex regulatory pattern in showing that (1) mtaCB1 and mtaCB2 are expressed much more highly (100- and 570-fold, respectively) on methanol than on acetate, that (2) mtaCB1 is expressed primarily during exponential growth phase while expression of mtaCB2 peaked in stationary growth phase, and that (3) expression of mtaCB3 is significantly up-regulated on MMA and acetate (Bose et al., 2006). From the data it was concluded that the three isogenes are regulated at multiple levels involving both positive and negative transcriptional regulation. This notion was corroborated by analyzing mutants, defective for cis-acting regulatory sequences in the vicinity of the mtaCB promoters, and for trans-acting proteins involved in mtaCB regulation (Bose and Metcalf, 2008). The genes encoding the regulators were designated msrA, msrB, msrC, msrD and msrE, respectively, which direct substrate-dependent expression of all three

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mtaCB operons in a concerted fashion. The study not only marks the first detailed investigation of the molecular mechanism underlying transcriptional regulation of any gene involved in methylotrophic methanogenesis, but also documents the first archaeal examples of regulatory proteins that simultaneously act in both repression and activation of transcription (Bose and Metcalf, 2008). Interestingly, growth on methanol under nitrogen limitation leads to increased expression of one of the MMA-specific MT1-encoding (mtmBC2) genes in M. mazei (Veit et al., 2005). Synthesis of a MMA-specific methyltransferase in the presence of methanol could be an adaptation strategy for a potential switch to an alternative energy source (Veit et al., 2005). Alternatively, it could also serve as a means to expand the range of potential nitrogen sources when this nutrient is limiting, as demethylation of MMA liberates ammonium. Such a strategy is employed by M. voltae upon selenium deprivation (Niess and Klein, 2004). The other methylamine-specific MT1 encoding genes of M. mazei are either up-regulated in the presence of TMA as compared to methanol (mtmBC1, mtbB1, mttBC1) or not affected (mtmBC1, mtbBC2, mttBC2). Expression profiling of the MT1 encoding genes on DMA and MMA was so far not reported, but would probably lead to similar results because utilization of TMA leads to DMA, and MMA formation (Hippe et al., 1979). Growth on methylated compounds leads to the induction of another important factor, F420H2 dehydrogenase (FPO, F420H2:methanophenazine oxidoreductase, Deppenmeier, 2004). Oxidation of methyl-groups from, e.g., methanol generates F420H2 (> Fig. 2), which is re-oxidized by FPO, transferring the electrons to the methanophenazine pool and two protons across the cytoplasmic membrane (Deppenmeier, 2002). The protein is encoded by the fpoABCDHIJKLMNO operon and is homologous to the NADH dehydrogenase from E. coli (Ba¨umer et al., 2000). As expected for an important coupling-site during methylotrophic methanogenesis, expression of the fpo genes is markedly increased (ca. tenfold) in M. mazei grown on methanol as compared to acetate (Hovey et al., 2005).

4

Acetotrophic Methanogenesis

The third pathway of methanogenesis is reviewed in > Chapter 23, Vol. 1, Part 5 (J. G. Ferry) and will therefore only briefly be summarized. Acetotrophic (aceticlastic) methanogenesis (Jetten et al., 1992, > Fig. 3) allows, compared to other substrates, only for very little energy to be conserved (36 kJ/mol, Mu¨ller et al., 1993). Consequently, genes not required for acetotrophic growth are generally strictly down-regulated (see above). Therefore, only the highly acetate-induced factors will be mentioned here (see > Table 1). Aceticlastic methanogenesis proceeds first by the activation of acetate to acetyl-CoA via phosphotransacetylase (Pta) and acetate kinase (Ack) (Ferry, 1997). The genes encoding the two proteins (pta and ack, respectively are commonly organized in an operon and their expression is highly induced on this substrate (Hovey et al., 2005; Li et al., 2006). An alternative enzyme for acetate activation, AMP-forming acetyl-CoA synthetase (AMP-ACS, encoded by acsA, Wolfe, 2005), is also up-regulated fourfold in M. mazei under aceticlastic conditions (Hovey et al., 2005). Subsequently, the bifunctional carbon monoxide dehydrogenase/acetyl-CoA synthase (CODH/ACS, also called ACDS) complex splits acetylCoA into enzyme-bound CO, a methyl-group, and free coenzyme A (Grahame, 1991). M. acetivorans, M. mazei and M. barkeri each encode two highly similar CODH/ACS isoforms (cdhABCDE 1 and 2)(Deppenmeier et al., 2002; Galagan et al., 2002; Maeder et al., 2006),

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whereas Methanosarcina thermophila contains only one cdh operon (Grahame et al., 2005). Expression of both cdh operons is highly induced in the former two organisms (7–14-fold higher than in methanol-grown cells, Hovey et al., 2005) leading to substantial amounts of CODH/ACS (10–250-fold higher than in methanol-grown cells, Li et al., 2006). CODH/ACS harbors a corrinoid-containing subunit, which donates the methyl-group to H4SPT. MethylH4SPT is reduced via reactions of the hydrogenotrophic pathway (> Fig. 3, and see above). The CO produced in the CODH/ACS-catalyzed reaction is oxidized to CO2 generating the electrons needed for reduction of the methyl-group to methane. The electron acceptor of CODH/ACS in methanogens has been shown to be ferredoxin (Fischer and Thauer, 1990). How the electrons are funneled to the terminal oxidase, heterodisulfide reductase, is not clear. In M. mazei, growth on acetate leads to an increase in mRNA abundance (3–10-fold) for genes encoding ECH (see above, Hovey et al., 2005), and a M. barkeri mutant lacking ECH is unable to grow with acetate (Meuer et al., 2002). These findings support the notion that H2 is an intermediate, and that this enzyme is involved in energy conservation, during aceticlastic methanogenesis of Methanosarcina (Meuer et al., 1999). In M. acetivorans, which lacks ECH, subunits of a Rnf-type membrane complex (Schmehl et al., 1993), homologous to NAD dehydrogenase from E. coli, are present at a tenfold higher level in acetate- than in methanol-grown cells (Li et al., 2006). It was speculated that ‘‘Rnf ’’ might function as a reduced ferredoxin:methanophenazine oxidoreductase and that it couples this exergonic reaction to ion translocation across the cytoplasmic membrane (Li et al., 2006), thus constituting a crucial component of the acetate-dependent respiratory chain in M. acetivorans. Evidence for this hypothesis, however, is currently unavailable.

5

Research Needs

Since the 1970s, when investigating the path of methane formation in ‘‘methanogenic bacteria’’ with biochemical and physiological analyses started, a wealth of novel knowledge about the unique nature of methanogenic metabolism was gained and the biochemistry of the paths for methane production is established today. The genomic information accumulating since the 1990s now allows to put that knowledge into the context of the genetic inventory and was a prerequisite to address global changes in mRNA- and protein-abundances under different conditions. Establishing methods for genetic analysis of methanogens added another angle for analyzing the metabolism and physiology of these unique and important prokaryotes. However, much can - and needs to - be learned about methanogens. Besides methanogenesis itself relatively little is known about other aspects of the methanoarchaea. As summarized here, genes involved in methanogenesis are regulated, but the mechanism of this regulation is largely undetermined. Furthermore, almost nothing is known about how changes in environmental conditions are sensed and how this information is processed. To solve these questions is an exciting task for the future, which will help to fully appreciate the global role methane and organisms producing it play.

Acknowledgments I am indebted to V. Mu¨ller, University of Frankfurt, for his generous support. I also thank W.W. Metcalf, University of Illinois, for stimulating discussions. Work in the authors’ laboratory is supported by grants from the Deutsche Forschungsgemeinschaft and the HerrmannWillkomm-Stiftung.

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Methanogenesis Purification, properties and encoding genes of the corrinoid protein MT1. Eur J Biochem 243: 670–677. Schmehl M, Jahn A, Meyer zu Vilsendorf A, Hennecke S, Masepohl B, Schuppler M, Marxer M, Oelze J, Klipp W (1993) Identification of a new class of nitrogen fixation genes in Rhodobacter capsulatus: a putative membrane complex involved in electron transport to nitrogenase. Mol Gen Genet 241: 602–615. Schmitz RA, Albracht SP, Thauer RK (1992) A molybdenum and a tungsten isoenzyme of formylmethanofuran dehydrogenase in the thermophilic archaeon Methanobacterium wolfei. Eur J Biochem 209: 1013–1018. Schwo¨rer B, Thauer RK (1991) Activities of formylmethanofuran dehydrogenase, methylenetetrahydromethanopterin dehydrogenase, methylenetetrahydromethanopterin reductase, and heterodisulfide reductase in methanogenic bacteria. Arch Microbiol 155: 459–465. Shima S, Lyon EJ, Sordel-Klippert M, Kauss M, Kahnt J, Thauer RK, Steinbach K, Xie X, Verdier L, Griesinger C (2004) The cofactor of the iron-sulfur cluster free hydrogenase hmd: structure of the lightinactivation product. Angew Chem Int Ed Eng 43: 2547–2251. Shima S, Thauer RK (2007) A third type of hydrogenase catalyzing H2 activation. Chem Rec 7: 37–46. Sorgenfrei O, Mu¨ller S, Pfeiffer M, Sniezko I, Klein A (1997) The [NiFe] hydrogenases of Methanococcus voltae: genes, enzymes and regulation. Arch Microbiol 167: 189–195. Steigerwald VJ, Hennigan AN, Pihl TD, Reeve JN (1993) Genes encoding the methyl viologen-reducing hydrogenase, polyferredoxin and methyl coenzyme M reductase II are adjacent in the genomes of Methanobacterium thermoautotrophicum and Methanothermus fervidus. In Microbial Growth on C1 Compounds. JC Murrell, DP Kelly (eds.). Andover, UK: Intercept Ltd., pp. 181–191. Sun J, Klein A (2004) A lysR-type regulator is involved in the negative regulation of genes encoding seleniumfree hydrogenases in the archaeon Methanococcus voltae. Mol Microbiol 52: 563–571.

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Tersteegen A, Hedderich R (1999) Methanobacterium thermoautotrophicum encodes two multisubunit membrane-bound [NiFe] hydrogenases. Eur J Biochem 264: 930–943. Thauer RK (1998) Biochemistry of methanogenesis: a tribute to Marjory Stephenson. Microbiology 144: 2377–2406. Thauer RK, Hedderich R, Fischer R (1993) Reactions and enzymes involved in methanogenesis from CO2 and H2. In Methanogenesis. JG Ferry (ed). New York: Chapman & Hall, pp. 209–252. Van Beelen P, Labro JFA, Keltjens JT, Geerts WJ, Vogels GD, Laarhoven WH, Guijt W, Haasnoot CAG (1984) Derivatives of methanopterin, a coenzyme involved in methanogenesis. Eur J Biochem 139: 359–366. Van der Maarel MJEC, Hansen TA (1997) Dimethylsulfoniopropionate in anoxic intertidal sediments: a precursor of methanogenesis via dimethyl sulfide, methanethiol, methiolpropionate. Mar Geol 137: 5–12. Veit K, Ehlers C, Schmitz RA (2005) Effects of nitrogen and carbon sources on transcription of soluble methyltransferases in Methanosarcina mazei strain Go¨1. J Bacteriol 187: 6147–6154. Vorholt JA, Vaupel M, Thauer RK (1997) A seleniumdependent and a selenium-independent formylmethanofuran dehydrogenase and their transcriptional regulation in the hyperthermophilic Methanopyrus kandleri. Mol Microbiol 23: 1033–1042. Wilting R, Schorling S, Persson BC, Bo¨ck A (1997) Selenoprotein synthesis in Archaea: identification of an mRNA element of Methanococcus jannaschii probably directing selenocysteine insertion. J Mol Biol 266: 637–641. Wolfe AJ (2005) The acetate switch. Microbiol Mol Biol Rev 69: 12–50. Zirngibl C, Van Dongen W, Schwo¨rer B, Von Bunau R, Richter M, Klein A, Thauer RK (1992) H2-forming methylenetetrahydromethanopterin dehydrogenase, a novel type of hydrogenase without iron-sulfur clusters in methanogenic archaea. Eur J Biochem 208: 511–520.

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37 Functional Genomics of Methanogens B. Lupa Department of Microbiology, The University of Georgia, Athens, GA, USA [email protected] 1 Functional Genomics – Definition, Goals, Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 502 2 Comparative Genomics of Methanogenic Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 502 3 Global Transcription and Proteomics Analyses – Applications for Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 503 4 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 507

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_37, # Springer-Verlag Berlin Heidelberg, 2010

502

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Functional Genomics of Methanogens

Abstract: Functional genomics is an interdisciplinary research field that focuses on determination of dynamic aspects of gene regulation, gene-gene, gene-protein and protein-protein interactions on the scale of the entire cell. Functional genomics is based on and integrates data from high throughput methods such as genomics, transcriptomics (microarrays), proteomics and bioinformatics. Thus, functional genomics provides the knowledge for the transition from the nucleotide sequence and protein function to complex cellular organization. Because of the increasing number of the sequenced genomes of methanogenic archaea, functional genomics studies are now feasible and are providing a significantly better understanding of the physiology of this important group of prokaryotes. In combination with genetic tools for generating various types of mutants, the influence of a given mutation on the gene expression and protein abundance on genome-wide scales can be examined by microarrays and proteomic studies. In addition, comparison of gene expression and protein abundance under various growth conditions, such as carbon, energy source, or nutrient limitations, have proven to be differential factors of great importance in understanding the physiology of methanogenic archaea. In the future, new experimental high throughput methods will be employed for methanogens. These approaches will facilitate a more comprehensive understanding of their physiology and complex interactions with other prokaryotes and animal hosts.

1

Functional Genomics – Definition, Goals, Methods

Functional genomics is an interdisciplinary research field that focuses on determination of function of genes and their products in the context of the entire genome or cell. The scope of functional genomics encompasses dynamic aspects of molecular biology and includes efforts for elucidation of regulation of gene expression, gene–gene interaction networks, and protein-protein interactions. Therefore, functional genomics provides a platform for the integration of experimental data from genomics, proteomics, protein crystallography and bioinformatics. The data matrices of levels of gene expression, ratios of protein abundances, information on post-translation modification, and concentrations of a large number of metabolites provide the basis for construction of predictive models of cell function, which constitutes the goal of systems biology (Ishii et al., 2004). This knowledge allows comprehensive understanding of the physiology of organisms, the cell itself, and its dynamic interactions with other cells and different elements of the surrounding environment. In the post-genomic era, the number and diversity of the revealed genomic sequences, particularly for microorganisms, allow for the transition from the nucleotide sequence to gene and protein function and finally to biological processes. Thus, functional genomics provides the basis for understanding the function of the gene products and their translation into the complex cell organization.

2

Comparative Genomics of Methanogenic Archaea

The genome is defined as the entire collection of DNA within an organism. By determination of the total nucleotide sequence, genome sequencing projects provide the insight into the physiology of organisms and are prerequisite for the functional genomics studies. The completed genomes of a number of methanogens facilitate the comparative studies (comparative genomics). Currently, 24 genomes of methanogenic archaea are sequenced, which

Functional Genomics of Methanogens

37

includes 18 validly published species, one candidatus and one reconstructed genome from the metagenomic project (> Table 1, for source of reference see Liu and Whitman, 2008). The genome lengths of methanogens vary from 1.57 to 5.75 million base pairs (Mbp). The genomes of Methanococcales and Methanobacteriales are typically smaller than 2 Mbp. In contrast, Methanosarcinales harbor the largest genomes, and the genome of Methanosarcina acetivorans is the largest yet revealed within Archaea. The G þ C content of the genomes of methanogens varies in a broad range of 27–61 mol %. The substantial portions of their genomes contain predicted open reading frames (ORFs) which encode the proteins with no assigned function. Therefore, revealing the functions of these genes and corresponding proteins became one of the main goals of functional genomics. Even though systems biology, including functional genomics, comprises the ultimate goal of obtaining the full knowledge of the biology of methanogens at the cellular, protein, gene, metabolite, and molecular level; the genome sequencing projects and comparative genomics provide valuable information about physiology and biochemistry of these microbes. The completed genome sequences provided information of the genetic basis for adaptations to the particular life style and relations between different groups of methanogens. For instance, the genome of the human gut inhabitant, Methanosphaera stadmanae revealed why this methanogen is restricted to utilization of methanol and H2 as substrates for methanogenesis and ATP synthesis (Fricke et al., 2006). The insights into mechanisms of thermal adaptations for two cold-adapted methanogens, Methangenium frigidum and Methanococcoides burtonii, were obtained from their genome sequences (Saunders et al., 2003). Likewise, the genome analysis of the hyperthermophilic Methanopyrus kandlerii provided the phylogenetic evidence for monophyletic origin of methanogenic archaea (Slesarev et al., 2002).

3

Global Transcription and Proteomics Analyses – Applications for Methanogens

Microarrays (DNA chips) are powerful tools for analysis of transcription in the whole-genome approach. The array contains a large number of the DNA samples and allows parallel investigation of the expression levels of thousands of genes within a cell. This method is based on the determination of the amount of mRNA which binds specifically to individual DNA oligonucleotides on the array. Subsequently, this information is used to construct the genome-wide gene expression profile in the cell. Typically, the investigated microorganism is cultivated under differential growth conditions followed by the microarrays analysis. In contrast, proteomics seeks to identify and quantify the relative abundances of proteins, on the scale of the entire cell, which may be differentially produced. The current progress of proteomics and its applications to prokaryotes, which includes the mesophile Methanococcus maripaludis, are discussed in a recent review (Xia et al., 2007). The global analysis of the ratio of protein abundances by its complex and dynamic nature is less developed than microarrays. In general, it involves the fractionation and separation of cellular proteins and their subsequent identification. Typically, the proteins are separated by two-dimensional polyacrylamide gel electrophoresis (2D-PAGE) and subjected to endopeptidase digestion. Alternatively, the mixture of proteins may be subjected to endopeptidase degradation, and the resulting peptides are separated by liquid chromatography. In either case, the peptides are identified by mass spectrometry.

503

504

37

Functional Genomics of Methanogens

. Table 1 Methanogenic archaea with completed genomic sequencing projectsa

Species/strain

Genome length (bp)

G+C Content (%)

Protein coding genes

RNA GenBank Acc. genes No.

Methanococci Methanococcus maripaludis S2

1,661,137

33

1722

50

BX950229

Methanococcus maripaludis C5

1,780,761

33

1813

47

CP000609

Methanococcus maripaludis C6

1,744,193

33

1826

48

CP000867

Methanococcus maripaludis C7

1,772,694

33

1788

47

CP000745

Methanococcus voltae A3

1,866,365

28

1690

36

ABHB00000000

Methanococcus voltae PSb

1,838,893

29

1700

Methanococcus aeolicus Nankai-3

1,569,500

30

1490

46

CP000743

Methanococcus vannielii SB

1,720,048

31

1678

51

CP000742

Methanocaldococcus jannaschii DSM 2661

1,664,970

31

1729

43

L77117

Methanothermococcus thermolithotrophicusb

1,731,708

33

1720

–

–

Methanomicrobia Methanocorpusculum labreanum Z

1,804,962

Methanospirillum hungatei JF-1

3,544,738

Methanoculleus marisnigri JR1 Candidatus Methanoregula boonei 6A8

1739

63

CP000559

45

3139

66

CP000254

2,478,101

62

2489

53

CP000562

2,542,943

54

2450

53

CP000780

Methanosaeta thermophila PT

1,879,471

53

1696

51

CP000477

Methanosarcina barkeri Fusaro

4,837,408

39

3606

73

CP000099

Methanosarcina mazei Go¨1

4,096,345

41

3370

66

AE008384

Methanosarcina acetivorans C2A

5,751,492

42

4540

69

AE010299

Methanococcoides burtonii DSM 6242

2,575,032

40

2273

63

CP000300

1,751,377

49

1873

48

AE000666

Methanosarcina

Methanobacteria Methanothermobacter thermautotrophicus Delta H

Functional Genomics of Methanogens

37

. Table 1 (Continued)

Species/strain

Genome length (bp)

G+C Content (%)

Protein coding genes

RNA GenBank Acc. genes No.

Methanobrevibacter smithii ATCC 35061

1,853,160

31

1793

42

CP000678

Methanosphaera stadtmanae DSM 3091

1,767,403

27

1534

52

CP000102

Uncultured methanogenic archaeon RC-1MRE50

3,179,916

54

3085

63

AM114193

Methanopyrus kandleri AV19 1,694,969

61

1687

38

AE009439

a

The partial genome sequences for an additional 32 methanogens are available in the Genomes OnLine Database, http://www.genomesonline.org b Data from ERGO database, Integrated Genomics

The microarray investigations of methanogenic archaea revealed important strategies of adaptation to different growth conditions and provided valuable insight into their metabolism. The global mRNA levels following H2, phosphate, and leucine limitation as well as growth at different growth rates was studied in M. maripaludis (Hendrickson et al., 2007, 2008). In these studies, cells were grown in chemostats with identical growth rates and cell densities. Differential expression was then compared between each of pair of conditions. These studies revealed the sets of functionally distinct genes which were specifically regulated by each limiting nutrient. The H2 limitation caused significant up-regulation of genes which encode proteins with the coenzyme F420-dependent functions. This included F420-reducing hydrogenase or Fru, formate dehydrogenase or Fdh, methylenetetrahydromethanoptherin dehydrogenase or Mtd and methylenetetrahydromethanompterin reductase or Mer. In addition, H2 limitation caused moderate increases in the mRNA abundance for genes with redox functions not involving F420, such as energy conserving hydrogenase b or Ehb, CO dehydrogenase-acetyl CoA synthase or CODH-ACS and pyruvate oxidoreductase or POR. Leucine limitation resulted in increased mRNA levels for a number of ribosomal proteins and RNA polymerase. In contrast, the mRNAs with functions in amino acid biosynthesis were only slightly affected. Phosphate limiting conditions caused significant up-regulation of mRNA for a phosphate ABC transporter (Hendrickson et al., 2008). Moreover, specific changes in mRNA levels for some genes could be assigned to growth rate effects. The gene hmd, which encodes the H2-dependent methylenetetrahyderomethanopterin dehydrogenase was up-regulated. In contrast, the genes encoding the heterodisulfide reductase or Hdr and F420-non reducing hydrogenase or Vhu were moderately down-regulated. In addition, increased growth rate caused increased levels of mRNA for ribosomal and S-layer proteins (Hendrickson et al., 2007, 2008). The microarray and proteomics analyses were used to study the function of the energyconserving, membrane-bound hydrogenase Ehb in M. maripaludis. This enzyme system is encoded by 16 genes scattered in 7 loci on the genome. It was hypothesized to couple H2 oxidation to the proton motive force for generation of a strong reductant for CO2 reduction. First, a mutant was constructed by replacement of ehbF, which encoded a large intrinsic membrane component of the complex, with the pac cassette to inactivate the complex.

505

506

37

Functional Genomics of Methanogens

The levels of mRNA were examined by microarray analysis and the proteome was examined by multidimensional liquid chromatography-mass spectrometry (LC MS/MS). The relative gene expression levels and ratios of protein abundance in the mutant were compared to the wild type strain S2. In this study, 55% of the proteins encoded by the genome were detected, and good correlation between the proteome and microarray data was observed. The mutant strain possessed higher levels for both mRNA and proteins which were involved in carbon assimilation. This included the CODH-ACS and POR. Both enzymes depend upon low potential ferredoxins for their biosynthetic activity, and changes in their expression were consistent with a specific role of Ehb in autotrophic CO2 assimilation instead of methanogenesis (Porat et al., 2006; Xia et al., 2006). The methylotrophic fresh water Methanosarcina mazei serves as the excellent model for genome-wide studies of differential expression levels in response to various growth conditions. M. mazei utilizes acetate, methanol, methylamines and H2 as substrates for methanogenesis and growth. The DNA-microarrays were used to study the transcription response to different methanogenic substrates. Acetate-grown cells expressed elevated levels of mRNA for genes encoding enzymes of the aceticlastic pathway that included CODH-ACS, carbonic anhydrase and acetate-activating enzymes, such as acetate kinase and phsophotransacetylase. Interestingly, the messages were also elevated for enzymes of energy metabolism, such as the membranebound hydrogenase Ech and A1A0-type ATP synthase. In addition, a number of genes were induced that encoded various ferredoxins, flavoproteins, aldehyde:ferredoxin oxidoreductase, enzymes of aromatic amino acid biosynthesis and proteins associated with uptake of cobalt, iron and oligonucleotides. The methanol-grown cells demonstrated the higher levels of mRNA for the enzymes of methanol disproportionation and translation (Hovey et al., 2005). Another study focused on the M. mazei stressed by high salt. These investigations revealed numbers of genes up-regulated by stress, including transporters for sodium and phosphate, stress response and regulatory proteins, DNA modification systems and cell surface modulators. In addition, the message for several hypothetical genes was highly (100–300 fold) elevated, which indicated a role in these conditions (Pfluger et al., 2007). Genome-wide transcription response to different N2 availability revealed number of genes that were up-regulated under the N2 limitation, which included genes with functions in N2 and carbon metabolism, stress response, transport as well as genes of unknown function (Veit et al., 2006). The transcriptomic and proteomic analyses of differentially grown marine M. acetivorans gave insight into the physiology of this versatile methanogen. The acetate-grown cells showed increased abundance of 255 proteins relative to methanol-grown cells. Likewise, the relative mRNA abundance was elevated for 410 genes in the acetate-grown cells. The expression levels from microarrays correlated well with the protein abundances under each condition. As a result, new information about the electron transport components specific for acetate-grown cells, the function of duplicate CODH-ACSs, general stress response and number of regulatory proteins was revealed. Similarly, over 200 genes were specifically up-regulated and nearly 190 proteins were detected as more abundant during the growth with methanol. This included the enzymes of the methanol-dependent methanogenesis and some anabolic pathways as well as genes involved in translation (Li et al., 2007). Another proteomics studies provided data about pathway of CO2 reduction to CH4 by CO-grown M. acetivorans. This pathway included novel methyltransferases and a new mechanism for energy conservation, the F420H2:CoM-S-S-CoB oxidoreductase. In addition, these studies revealed the increased abundance of the monofunctional CODH during the CO-dependent growth (Lessner et al., 2006; Rother et al., 2007).

Functional Genomics of Methanogens

37

In contrast to the relatively well studied mesophilic species, the proteomic analyses of extremophilic methanogens are scarcer. The global study of protein abundance ratios of the hyperthermophilic M. jannaschii revealed that the most abundant cellular proteins produced under optimal growth conditions have no assigned function (Giometti et al., 2002). Likewise, insight into physiology and mechanisms of adaptation to low temperature of the psychrophilic methanogenic archaeon, Methanococcoides burtonii, was obtained by the proteomics studies (Goodchild et al., 2004). This study revealed expression of transposonase and a number of other proteins that may be specifically induced during growth at 4  C.

4

Research Needs

Despite rapid progress in development of functional genomics and its application to Archaea and in particular to methanogens, the field remains open for new improvements and the implementation of new research techniques. This is required for the progress in understanding the genetics, biochemistry and lifestyle of methanogenic archaea. In the near future, the elucidation of functions of the predicted, hypothetical genes that encode proteins with unknown functions is of great importance. Current limitations of high throughput methods, especially proteomics should be addressed. The peptide separation methods, signal to noise ratio, and unambiguous peptide identification must be improved to overcome the problems facing quantification of differential expression on a global scale. Progress in comparative genomics is necessary to fully evaluate the additional genomes of methanogenic archaea which have been or are currently being sequenced (see Genomes OnLine Database, http://www.genomesonline.org). Additional genomic sequences are required to fully explore groups of methanogens, such as the family Methanothermaceae or the unclassified genus Methanocalculus. The new research tools such as antibody arrays and whole proteome chips could be useful for analysis of protein-protein interactions. The global analysis of protein functions in vivo could be assayed by cell arrays. Nanoarrays could provide the knowledge of physiology of methanogens at the molecular level. Bioinformatics should provide novel tools to allow the analysis of large sets of data that are generated in new high throughput methods. Finally, the integration of genomics, proteomics, and metabolomics data will drive the progress of systems biology, an increased understanding of the methanogen diversity in the environment, and an increased capacity to utilize these microbes in biotechnological applications.

Acknowledgments The author expresses his gratitude to Dr. William B. Whitman for the critical reading of the manuscript and helpful comments.

References Fricke WF, Seedorf H, Henne A, Kruer M, Liesegang H, Hedderich R, Gottschalk G, Thauer RK (2006) The genome sequence of Methanosphaera stadtmanae

reveals why this human intestinal archaeon is restricted to methanol and H2 for methane formation and ATP synthesis. J Bacteriol 188: 642–658.

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Giometti CS, Reich C, Tollaksen S, Babnigg G, Lim H, Zhu W, Yates J, Olsen G (2002) Global analysis of a ‘‘simple’’ proteome: Methanococcus jannaschii. J Chromatogr B Analyt Technol Biomed Life Sci 782: 227–243. Goodchild A, Raftery M, Saunders NF, Guilhaus M, Cavicchioli R (2004) Biology of the cold adapted archaeon, Methanococcoides burtonii determined by proteomics using liquid chromatography-tandem mass spectrometry. J Proteome Res 3: 1164–1176. Hendrickson EL, Haydock AK, Moore BC, Whitman WB, Leigh JA (2007) Functionally distinct genes regulated by hydrogen limitation and growth rate in methanogenic Archaea. Proc Natl Acad Sci USA 104: 8930–8934. Hendrickson EL, Liu Y, Rosas-Sandoval G, Porat I, Soll D, Whitman WB, Leigh JA (2008) Global Responses of Methanococcus maripaludis to Specific Nutrient Limitations and Growth Rate. J Bacteriol 190: 2198–2205. Hovey R, Lentes S, Ehrenreich A, Salmon K, Saba K, Gottschalk G, Gunsalus RP, Deppenmeier U (2005) DNA microarray analysis of Methanosarcina mazei Go1 reveals adaptation to different methanogenic substrates. Mol Genet Genomics 273: 225–239. Ishii N, Robert M, Nakayama Y, Kanai A, Tomita M (2004) Toward large-scale modeling of the microbial cell for computer simulation. J Biotechnol 113: 281–294. Lessner DJ, Li L, Li Q, Rejtar T, Andreev VP, Reichlen M, Hill K, Moran JJ, Karger BL, Ferry JG (2006) An unconventional pathway for reduction of CO2 to methane in CO-grown Methanosarcina acetivorans revealed by proteomics. Proc Natl Acad Sci USA 103: 17921–17926. Li L, Li Q, Rohlin L, Kim U, Salmon K, Rejtar T, Gunsalus RP, Karger BL, Ferry JG (2007) Quantitative proteomic and microarray analysis of the archaeon Methanosarcina acetivorans grown with acetate versus methanol. J Proteome Res 6: 759–771. Liu Y, Whitman WB (2008) Metabolic, phylogenetic, and ecological diversity of the methanogenic archaea. Ann N Y Acad Sci 1125: 171–189. Pfluger K, Ehrenreich A, Salmon K, Gunsalus RP, Deppenmeier U, Gottschalk G, Muller V (2007) Identification of genes involved in salt adaptation

in the archaeon Methanosarcina mazei Go1 using genome-wide gene expression profiling. FEMS Microbiol Lett 277: 79–89. Porat I, Kim W, Hendrickson EL, Xia Q, Zhang Y, Wang T, Taub F, Moore BC, Anderson IJ, Hackett M, Leigh JA, Whitman WB (2006) Disruption of the operon encoding Ehb hydrogenase limits anabolic CO2 assimilation in the archaeon Methanococcus maripaludis. J Bacteriol 188: 1373–1380. Rother M, Oelgeschlager E, Metcalf WM (2007) Genetic and proteomic analyses of CO utilization by Methanosarcina acetivorans. Arch Microbiol 188: 463–472. Saunders NF, Thomas T, Curmi PM, Mattick JS, Kuczek E, Slade R, Davis J, Franzmann PD, Boone D, Rusterholtz K, Feldman R, Gates C, Bench S, Sowers K, Kadner K, Aerts A, Dehal P, Detter C, Glavina T, Lucas S, Richardson P, Larimer F, Hauser L, Land M, Cavicchioli R (2003) Mechanisms of thermal adaptation revealed from the genomes of the Antarctic Archaea Methanogenium frigidum and Methanococcoides burtonii. Genome Res 13: 1580–1588. Slesarev AI, Mezhevaya KV, Makarova KS, Polushin NN, Shcherbinina OV, Shakhova VV, Belova GI, Aravind L, Natale DA, Rogozin IB, Tatusov RL, Wolf YI, Stetter KO, Malykh AG, Koonin EV, Kozyavkin SA (2002) The complete genome of hyperthermophile Methanopyrus kandleri AV19 and monophyly of archaeal methanogens. Proc Natl Acad Sci USA 99: 4644–4649. Veit K, Ehlers C, Ehrenreich A, Salmon K, Hovey R, Gunsalus RP, Deppenmeier U, Schmitz RA (2006) Global transcriptional analysis of Methanosarcina mazei strain Go1 under different nitrogen availabilities. Mol Genet Genomics 276: 41–55. Xia Q, Hendrickson EL, Wang T, Lamont RJ, Leigh JA, Hackett M (2007) Protein abundance ratios for global studies of prokaryotes. Proteomics 7: 2904–2919. Xia Q, Hendrickson EL, Zhang Y, Wang T, Taub F, Moore BC, Porat I, Whitman WB, Hackett M, Leigh JA (2006) Quantitative proteomics of the archaeon Methanococcus maripaludis validated by microarray analysis and real time PCR. Mol Cell Proteomics 5: 868–881.

38 Regulation of Membrane Lipid Homeostasis in Bacteria M. A. Martinez . G. E. Schujman . H. C. Gramajo . D. de Mendoza* Instituto de Biologı´a Molecular y Celular de Rosario (IBR-CONICET), Departamento de Microbiologı´a, Facultad de Ciencias Bioquı´micas y Farmace´uticas, Universidad Nacional de Rosario, Suipacha 531, Rosario, Argentina *[email protected] [email protected] [email protected] [email protected] 1 Introduction: Overview of the Reactions of Fatty Acid Biosynthesis . . . . . . . . . . . . . . . 510 2 Biosynthesis of UFA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 510 3 Control of Fatty Acid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 514 4 Biochemical Control of Fatty Acid Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 514 5 Transcriptional Control of FASII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 515 6 Control of Membrane Fluidity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 515 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 516

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_38, # Springer-Verlag Berlin Heidelberg, 2010

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38

Regulation of Membrane Lipid Homeostasis in Bacteria

Abstract: Fatty acid biosynthesis is an essential process for bacteria, but it is also energetically expensive. In addition, bacterial survival depends on membrane lipid homeostasis and on the ability to adjust lipid composition to acclimatize the bacterial cells to several environments. Thus, bacteria have developed homeostatic mechanisms that maintain the concentration of lipids at particular levels. Fatty acid synthesis in bacteria is achieved by a highly conserved set of genes in which each gene encodes an individual enzymatic step in the type II biosynthetic pathway. All of the proteins in this pathway are located in the cytosol, and each has been purified and biochemically characterized. Furthermore, the structure and catalytic mechanism of each enzyme is known in considerable detail. Although bacteria precisely and stringently control the synthesis of their membrane lipids, the regulatory mechanisms are incompletely understood. Here, the principal genetic and biochemical processes that are responsible for membrane lipid homeostasis are reviewed.

1

Introduction: Overview of the Reactions of Fatty Acid Biosynthesis

The first committed step of fatty acid biosynthesis is the conversion of acetyl-CoA to malonylCoA catalyzed by the enzyme acetyl-CoA carboxylase (ACC) (> Fig. 1). Malonyl-CoA is then used by the Type II or dissociated fatty acid enzymes to make long-chain fatty acids via a repeated cycle of reactions involving the condensation, reduction, dehydration and reduction of carbon-carbon bonds (de Mendoza et al., 2002; Lu et al., 2004). A key feature of this system is that all fatty acyl intermediates are covalently connected with a small acidic protein, acyl carrier protein (ACP), and shuttled from one enzyme to another until the acyl-ACPs reach the proper length and become substrates for the acyltransferases that transfer the fatty acyl chain into the membrane phospholipids. This biosynthetic scheme is conserved in all fatty acid producing bacteria, but substrate specificity of some of the enzymes involved in the pathway leads to the variety of fatty acids found in the different bacterial genera (Campbell and Cronan, 2001; Lu et al., 2004). Different bacteria have evolved distinct mechanisms for unsaturated fatty acid biosynthesis and regulation (reviewed in Lu et al., 2004; Schujman and de Mendoza, 2005, 2008).

2

Biosynthesis of UFA

Bacterial growth requires a considerable fraction of the acyl chains of the membrane lipids to be in a disordered state (‘‘fluid’’). A disordered state is imparted by presence of either cisunsaturated or terminally anteiso-branched-chain fatty acids; both of which act to offset the closely packed ordered arrangement imparted by the straight saturated acyl chains of the lipid bilayer. In this section, we discuss the basic features of UFA biosynthesis in organisms in which the proposed biosynthetic mechanisms have been characterized. In E. coli, synthesis of the normal UFA content requires three enzymes, FabA, FabB and FabF (> Fig. 1). FabA, a b-hydroxydecanoyl-ACP dehydrase, which normally elongates the C10 intermediate to C12 and beyond to produce saturated fatty acid, is also the key enzyme of the classic anaerobic pathway of UFA synthesis by introducing a cis double bond into a 10-carbon intermediate. The property of FabA that account for this bi-functional activity lies in that this enzyme is able to catalyzes both the removal of water to generate trans-2-decenoyl-ACP and

. Figure 1 Fatty acid metabolism in bacteria and its regulation. The initiation and elongation steps of type II fatty acid biosynthesis are boxed in light gray. The UFA biosynthesis in B. subtilis, P. aeruginosa and E. coli is boxed in dark gray. Regulatory circuits of B. subtilis are indicated with short-dashed lines; of E.coli with long-dashed lines and of P. aeruginosa with pointed lines. Activation circuits are shown with an arrow ending line (!), and repression circuits with a bar ending line (┬). Transcriptional regulators are enclosed in a double rectangle and metabolic effectors are depicted with a pointed-line rectangle. For clarity, general reactions are depicted. B. subtilis contains two isoforms of FabH and no FabB. PlsY acyltransferase is not under transcriptional control of FapR (for details see text).

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the isomerization of this intermediate to cis-3-decenoyl-ACP. This cis-10 carbon intermediate is then elongated by FabB, a 3-oxoacyl-ACP synthase I and later by FabF (3-oxoacyl-ACP synthase II), to form the two major UFA found in E. coli: C16:1D9 and C18:1D11. FabB and FabF have distinct and non-overlapping roles in E. coli UFA synthesis. FabB is thought to elongate the product of the FabA enzyme to the C12 unsaturated intermediate, whereas FabF is required to convert C16:1 unsaturated species to the C18:1 species (> Fig. 1). Expression of the key UFA synthetic gene fabA from E. coli is positively regulated by FadR which was hitherto thought to function only as a repressor of the ß-oxidation pathway of fatty acids (Henry and Cronan, 1992; Iram and Cronan, 2005). Positive regulation of fabA expression is neutralized both, in vivo and in vitro, by long-chain acyl-CoAs that are small ligands which regulate the binding of FadR to the fabA promoter. FadR is also a positive regulator of fabB, the second UFA biosynthetic gene, although the changes in fabB expression in fadR mutants are not as large as with fabA. Thus, FadR acts as a repressor of ß-oxidation genes and as an activator of the two genes required for UFA synthesis. FabR is a second E. coli regulator of UFAs synthesis (Zhang et al., 2002). It behaves as a repressor acting downstream of the FadR operator sequences in the regulation of fabA and fabB. The ligand regulating FabR activity is still unknown. There is a variation of the FabA/FabB bacterial strategy for UFA synthesis, discovered in Enterococcus faecalis. This organism possesses a special (3R)-hydroxymyristoyl-ACP dehydrase called FabN that also has a FabA like activity, and a unique FabF like enzyme, called FabO that replaces FabB. However, other organisms, like S. pneumoniae compensate FabA absence with an enzyme called FabM. This enzyme is capable of isomerizing the trans-unsaturated bond at the key 10-carbon intermediate, to its cis-isomer. In the examples mentioned above, the proteins FabA, FabM or FabN introduce the double bond into a C10 intermediate via an anaerobic reaction and either FabB or FabF channel this intermediate to the mainstream of the fatty acid synthetic pathway. In contrast, other bacteria like Bacillus and Cyanobacteria have completely separate systems for the synthesis of UFA and saturated fatty acids. These organisms use fatty acid desaturase enzymes, which require molecular oxygen and reducing equivalents, obtained from an electron transport chain, to introduce the double bond into previously synthesized saturated fatty acids (> Fig. 2). The organisms mentioned above each have only a single pathway for the synthesis of the UFAs required to make functional membrane lipids. In marked contrast, UFA biosynthesis in Pseudomonas aureginosa, proceeds by three distinct pathways. In this organism the FabA-FabB pathway does the bulk of UFA synthesis under all growth conditions. However, P. aeuroginosa has two fatty acyl desaturases (DesA and DesB) that supplement the FabA-FabB pathway under aerobic conditions and thereby allows faster growth. DesA introduces the double bond into the acyl chains of intact phospholipids, whereas the substrates of DesB are acyl-CoAs that are derived of exogenous fatty acids. DesB is inducible and is regulated by DesT, a transcriptional regulator that has the property of being able to sense the fatty acid composition of the long chain acyl-CoA pools to adjust the expression of the desaturase. Streptomyces, a soil microorganism that produces a vast array of pharmacologically important secondary metabolites, also adapts the content of its fatty acids in response to changes in temperature (Suutari and Laakso, 1992). Although these microorganisms have been less studied, they also appear to have more than one biosynthetic pathway for UFAs. Streptomyces coelicolor, for example, possess five fatty acid desaturases encoded on its genome, and some of them appear to be regulated either at the transcriptional or at the

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. Figure 2 Model pathways leading to membrane fluidity optimization. (a) In B. subtilis, a kinase dominant state of DesK predominates upon an increase in the proportion of ordered membrane lipids (due to temperature downshift). The phosphate group is transferred to the Asp54 of DesR, which in turn, activates the transcription of the des gene coding for the D5-desaturase (Des). Desaturation of membrane phospholipid acyl chains decreases membrane lipid order favoring the phosphatase-dominant state of DesK. Dephosphorylation of DesR results in decreased transcription of the des gene. (b) In E. coli, FabF participates in SFA synthesis and in the elongation of C16:1D9 palmitoleoyl-ACP to C18:1D11 cis-vaccenoyl-ACP. The reactivity of this enzyme toward C16:1D9 is increased after a temperature downshift, what accounts for the greater proportion of C18:1 acyl chains in phospholipids of bacteria growing at low temperatures compared to higher temperatures.

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translational level (Cole et al., 1998). The SCO3128 desaturase, for example, is forming an operon with a transcriptional regulator of the TetR family and the SCO3758 is also arranged in an operon fashion with a two component system of response regulators. The SCO6717, on the other hand, contains a TTA codon which is a well-known target for the translational regulation through the BldA tRNA (Gramajo et al., 1993). Besides the presence of five desaturases, Streptomyces also appear to generate UFAs through the anaerobic pathway, since they seem to be extremely sensitive to the specific dehydrase inhibitor 3-DecenoylNAC. On this regard, in silico analysis of several Streptomyces genomes revealed that they all have a putative enoyl-CoA isomerase highly similar to FabM, although no FabA or FabB homologues could be found.

3

Control of Fatty Acid Biosynthesis

Fatty acid synthesis is coordinately regulated with phospholipid, macromolecular synthesis and growth as part of the response to changes in the environment. This control is crucial for membrane homeostasis, because the biophysical properties of membranes are determined in large part by the composition that results from de novo lipid biosynthesis. Many of these processes are rapid responses of the integrated biochemical network and do not involve changes in gene expression. An important recent development is the identification and characterization of transcription factors that globally coordinate the expression level of genes that function in lipid biosynthesis.

4

Biochemical Control of Fatty Acid Biosynthesis

Elegant genetic experiments have demonstrated that the ACC enzymatic complex is the key component in determining the rate of fatty acid biosynthesis in E. coli, although not the only one. For instance, overexpression of the four acc genes in E. coli results in only sixfold increase in the rate of fatty acid synthesis while the malonyl-CoA level increases over 100-fold (Davis et al., 2000), indicating that later steps in the pathway also regulate the flux through the fatty acid synthetic pathway. The metabolic regulation of ACC occurs through a strong inhibition of the enzimatic activity by medium and long-chain acyl-ACPs (C6–C20), in a manner that is neither purely competitive nor noncompetitive with respect to acetyl-CoA (Davis and Cronan, 2001). Besides ACC, other enzymes of the FASII cycle, like FabH, which catalyzes the first step of the pathway and FabI, which catalyzes the completion of acyl-ACPs elongation (> Fig. 1) are also inhibited in vitro by long-chain acyl-ACPs (Heath and Rock, 1996). Recent evidences showed that PlsX (acyl-acyl carrier protein [ACP]:phosphate acyltransferase), the peripheral enzyme that catalyzes the first committed step in the biosynthesis of phospholipids in B. subtilis, plays an important role in the coordination of the production of fatty acids and phospholipids (Paoletti et al., 2007). In this organism the inactivation of PlsX leads to the cessation of both fatty acid and phospholipid synthesis. The inhibition of total lipid synthesis in PlsX-depleted cells was attributed to the accumulation of acyl-ACPs due to the block in its conversion to acyl-phosphate. This observation indicates that acyl-ACPs also downregulate fatty acid biosynthesis in B. subtilis.

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Transcriptional Control of FASII

A major advance in our understanding of the transcriptional control of bacterial lipid synthesis occurred following the identification of FapR, a global transcriptional repressor that controls the expression of the fap regulon, involved in the biosynthesis of fatty acids and phospholipids in B. subtilis (Schujman et al., 2003). The binding of FapR to its DNA targets is specifically inhibited by malonyl-CoA, an essential intermediate of fatty acid synthesis in all living cells, the cellular pools of which provide a mechanism for sensing the status of fatty acid biosynthesis and for adjusting the fap regulon accordingly (Schujman et al., 2006). FapR is highly conserved in many Gram-positive organisms, many of which are human pathogens, including Bacillus anthracis, Bacillus cereus, Listeria monocytogenes and Staphylococcus aureus. The consensus binding sequence of FapR is also highly conserved in the putative promoter regions of the fapR gene in those species. These observations indicate that the regulation mechanism observed in B. subtilis is probably conserved in many other organisms. The recent crystal structure of the effector binding domain of FapR, alone and complexed with malonyl-CoA, has provided the first insight into the ligand-induced transcriptional regulation of fatty acid and phospholipid synthesis (Schujman et al., 2006). FapR is a homodimeric protein that displays the typical ‘‘hot-dog’’ fold characteristic of the thioesterase enzyme family. Binding of malonyl-CoA promotes an order-disorder transition, which transforms an open ligand-binding groove into a long tunnel occupied by the effector molecule in the complex. This ligand-induced modification propagates to the helix-turn-helix motive, impairing their productive association for DNA binding. Structure-based mutations that disrupt the FapR-malonyl-CoA interaction prevent DNA binding regulation and results in a lethal phenotype in B. subtilis, suggesting this homeostatic signaling pathway as a promising target for novel chemotherapeutic agents against Gram-positive pathogens. The second global regulator, FabT, was identified in the human pathogen S. pneumoniae (Lu and Rock, 2006). In this bacterium, the twelve genes required for fatty acid biosynthesis are located together in a single cluster. The second gene in the cluster encodes the FabT transcriptional repressor, which regulates all these genes except fabM (involved in UFA biosynthesis). A similar genetic organization is found in other Gram-positive bacteria, such as Enterococcus, Clostridium and Lactococcus. The ligand that modulates the binding of FabT to its DNA targets is still unknown. In the actinomycete S. coelicolor a new class of transcriptional regulator of the fatty acid biosynthesis genes has been recently found and is being characterized (Arabolaza et al., unpublished). Interestingly, this transcriptional regulator is not essential for the bacterial survival, although mutants in it are severely affected on their growth capacity. Another important characteristic of this system is that the regulatory protein behaves as a transcriptional activator, in contrast with most of the previously reported regulators of fatty acid biosynthesis that negatively regulate the expression of fas genes. The regulator does not present a significant homology with anyone of the previously characterized transcriptional regulators in bacteria.

6

Control of Membrane Fluidity

Bacteria can encounter a wide range of environments and must adapt to new conditions in order to survive. As the selective barrier between living cells and their environment, the plasma

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membrane plays a key role in cell viability. The barrier function of the cytoplasmic membrane is known to depend critically on the physical state of lipid bilayers, making it susceptible to changes in environmental temperature. In fact, it has been established that lipid bilayers of most organisms are entirely or mostly fluid at physiological temperatures allowing normal cell functions. However, at lower temperatures, membrane lipid bilayers undergo a reversible change of state from a fluid (disordered) to a nonfluid (ordered) array of the fatty acyl chains. Therefore, bacteria must regulate their membrane fluidity in response to temperature in order to restore its normal cellular function. Without regulation, an organism shifted from a high to a low temperature would have membrane lipids with suboptimal fluidity, resulting in subnormal membrane function. The mechanism of regulation in all of the cases examined seems to occur via the incorporation of proportionally more UFAs (or fatty acids of analogous properties) as the temperature decreases. This regulatory mechanism, called thermal control of fatty acid synthesis, seems to be a universally conserved adaptation response allowing cells to maintain the appropriate fluidity of membrane lipids regardless of the ambient temperature. In this section, we discuss the basic features of thermal regulation of membrane lipid fluidity in E. coli and B. subtilis, in which the proposed mechanisms are firmly based on both genetic and biochemical evidence. There are two fundamental differences between the E. coli and B. subtilis models that account for the formation of UFAs and the regulation by growth temperature of the synthesis of these fatty acids. First, B. subtilis lacks the fabB and fabA genes that are essential for UFA synthesis in E. coli, and this property accounts for the absence of UFA production by the Gram positive synthase. Instead, B. subtilis makes UFAs via an oxygen-dependent desaturase that uses existing membrane phospholipids as substrates to introduce a double bond at the fifth position of the fatty acyl chain (Aguilar et al., 2001). Second B. subtilis uses a two-component system composed of a membrane-associated kinase, DesK, and a transcriptional regulator, DesR, which stringently control the transcription of the gene coding for the desaturase. Induction of this pathway under a temperature downshift is brought about via the ability of the molecular thermosensor DesK to sense a decrease in membrane fluidity (> Fig. 2). In contrast, in E. coli there is a biochemical mechanism that allows the elongation cycle of fatty acid synthesis to respond to the environmental temperature. In this organism the UFA synthesized in greater quantity at low temperatures is C18:1D11. This regulatory response is due to the properties of FabF that converts C16:1D9–C18:1D11. This enzyme is present at all temperatures but is more active at low temperature (> Fig. 2). Thus, a cytosolic thermosensor governs the temperature-dependent adjustment on membrane fluidity in E. coli, and probably in related bacteria, while in B. subtilis this control is exerted by a membrane-associated thermosensor.

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Research Needs

Desaturase expression is governed by complex signaling pathways, involving sensors that can adopt alternative states that are regulated by the biophysical properties of the membrane and coupled to transcription factors that control lipid homeostasis. An interesting unsolved question is how the sensors of membrane fluidity operate at the molecular level. Another relevant unanswered question related to transcription factors regulating lipid biosynthesis in Gram positive bacteria is how the level of the signal molecule modulating FapR activity varies in different growth and environmental conditions. In E. coli, malonyl-CoA levels are mainly the result of the competition between the activities of the biosynthetic ACC

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complex and the FabF condensing enzyme. Little is known in the paradigmatic bacterium B. subtilis about the factors that control ACC activity or the biochemical properties of FabF.

References Aguilar PS, Hernandez-Arriaga AM, Cybulski LE, Erazo AC, de MD (2001) Molecular basis of thermosensing: a two-component signal transduction thermometer in Bacillus subtilis. EMBO J 20: 1681–1691. Campbell JW, Cronan JE Jr (2001) Bacterial fatty acid biosynthesis: targets for antibacterial drug discovery. Annu Rev Microbiol 55: 305–332. Cole ST, Brosch R, Parkhill J, Garnier T, Churcher C, Harris D, et al. (1998) Deciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature 393: 537–544. Davis MS, Cronan JE Jr (2001) Inhibition of Escherichia coli acetyl coenzyme A carboxylase by acyl-acyl carrier protein. J Bacteriol 183: 1499–1503. Davis MS, Solbiati J, Cronan JE Jr (2000) Overproduction of acetyl-CoA carboxylase activity increases the rate of fatty acid biosynthesis in Escherichia coli. J Biol Chem 275: 28593–28598. de Mendoza D, Schujman GE, Aguilar PS (2002) Biosynthesis and function of membrane lipids. In Bacillus subtilis and its Closest Relatives: From Genes to Cells. AL Sonenshein, JA Hoch, and R Losick (eds.). Washington, DC: ASM Press, pp. 43–55. Gramajo HC, Takano E, Bibb MJ (1993) Stationaryphase production of the antibiotic actinorhodin in Streptomyces coelicolor A3(2) is transcriptionally regulated. Mol Microbiol 7: 837–845. Heath RJ, Rock CO (1996) Regulation of fatty acid elongation and initiation by acyl-acyl carrier protein in Escherichia coli. J Biol Chem 271: 1833–1836. Henry MF, Cronan JE Jr (1992) A new mechanism of transcriptional regulation: release of an activator triggered by small molecule binding. Cell 70: 671–679.

Iram SH, Cronan JE (2005) Unexpected functional diversity among FadR fatty acid transcriptional regulatory proteins. J Biol Chem 280: 32148–32156. Lu YJ, Rock CO (2006) Transcriptional regulation of fatty acid biosynthesis in Streptococcus pneumoniae. Mol Microbiol 59: 551–566. Lu YJ, Zhang YM, Rock CO (2004) Product diversity and regulation of type II fatty acid synthases. Biochem Cell Biol 82: 145–155. Paoletti L, Lu YJ, Schujman GE, de MD, Rock CO (2007) Coupling of fatty acid and phospholipid synthesis in Bacillus subtilis. J Bacteriol 189: 5816–5824. Schujman GE, de Mendoza D (2005) Transcriptional control of membrane lipid synthesis in bacteria. Curr Opin Microbiol 8: 149–153. Schujman GE, de Mendoza D (2008) Regulation of type II fatty acid synthase in Gram-positive bacteria. Curr Opin Microbiol 11: 148–152. Schujman GE, Guerin M, Buschiazzo A, Schaeffer F, Llarrull LI, Reh G, et al. (2006) Structural basis of lipid biosynthesis regulation in Gram-positive bacteria. EMBO J 25: 4074–4083. Schujman GE, Paoletti L, Grossman AD, de Mendoza D (2003) FapR, a bacterial transcription factor involved in global regulation of membrane lipid biosynthesis. Dev Cell 4: 663–672. Suutari M, Laakso S (1992) Changes in fatty acid branching and unsaturation of Streptomyces griseus and Brevibacterium fermentans as a response to growth temperature. Appl Environ Microbiol 58: 2338–2340. Zhang YM, Marrakchi H, Rock CO (2002) The FabR (YijC) transcription factor regulates unsaturated fatty acid biosynthesis in Escherichia coli. J Biol Chem 277: 15558–15565.

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39 Type III Polyketide Synthases Responsible for Phenolic Lipid Synthesis A. Miyanaga . S. Horinouchi* Department of Biotechnology, Graduate School of Agriculture and Life Sciences, University of Tokyo, Bunkyo-ku, Tokyo, Japan *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 520 2 The ars Operon in Azotobacter vinelandii . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 521 3 The srs Operon in Streptomyces griseus . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 522 4 Other Type III PKSs Responsible for Phenolic Lipid Synthesis . . . . . . . . . . . . . . . . . . . . . 523 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 524

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_39, # Springer-Verlag Berlin Heidelberg, 2010

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Type III Polyketide Synthases Responsible for Phenolic Lipid Synthesis

Abstract: In various microorganisms, phenolic lipids, consisting of polar aromatic rings and hydrophobic alkyl chains, are synthesized by type III polyketide synthases (PKSs). In Azotobacter vinelandii, two type III PKSs, ArsB and ArsC, are responsible for the biosynthesis of alkylresorcinols and alkylpyrones, which are essential for cyst formation. In Streptomyces griseus, a type III PKS, SrsA, is involved in the biosynthesis of alkylresorcinols, which confer resistance to b-lactam antibiotics on the host. Ars- and Srs-type PKSs are distributed not only in a wide variety of Gram-positive and negative bacteria but also in filamentous fungi. The phenolic lipids produced by type III PKSs thus play an important, but thus far overlooked, role as minor components in the biological membrane in both prokaryotes and eukaryotes.

1

Introduction

Polyketides form a large family of natural products found in bacteria, fungi and plants, and show various biological activities. Polyketides are synthesized by polyketide synthases (PKSs). PKSs possess a ketosynthase activity that catalyzes the sequential head-to-tail incorporation of two-carbon acetate units from a malonate thioester into a growing polyketide chain. Type III PKSs, consisting of a homodimeric ketosynthase, are the simplest enzymes among three types of PKSs, types I to III (Austin and Noel, 2003). Formerly, it had been thought that type III PKSs were distributed exclusively in plants. Many plant type III PKSs were discovered and characterized. For example, chalcone synthase catalyzes the formation of naringenin chalcone, an essential precursor of flavonoids and isoflavonoids, which have attracted significant attention because of their wide range of pharmacological properties. The first bacterial type III PKS, RppA, was reported in 1999 (Funa et al., 1999). RppA converts five malonyl-CoA molecules into 1,3,6,8-tetrahydroxynaphthalene, which serves as an intermediate in the biosynthesis of melanin and secondary metabolites in the actinobacterium, Streptomyces griseus. PhlD, a type III PKS in Pseudomonas fluorescens, is responsible for the biosynthesis of 2,4-diacetylphloroglucinol with biocontrol activity against soilborne fungal plant pathogens (Bangera and Thomashow, 1999). DpgA, a type III PKS from the actinobacterium Amycolatopsis mediterranei, is involved in the biosynthesis of glycopeptides antibiotics such as vancomycin (Pfeifer et al., 2001). The discoveries of these type III PKSs have established the idea that type III PKSs are distributed widely in microorganisms. The genome sequences of many organisms in the databases bioinformatically predict that type III PKSs are distributed widely in microorganism. On the basis of the bioinformatic prediction and subsequent in vivo and in vitro analyses, a number of type III PKSs have been identified to be responsible for the biosynthesis of secondary metabolites, such as antibiotics and pigments. For example, steely, a fusion enzyme of a type I FAS-type III PKS, from a social amoeba Dictyostelium discoidem, synthesizes a differentiation-inducing factor (Austin et al., 2006). In addition, some examples of type III PKSs involved in the biosynthesis of phenolic lipids such as alkylresorcinols and alkylpyrones were reported. Alkylresorcinols are distributed widely in bacteria, fungi and plants (Kozubek and Tyman, 1999). They consist of polar aromatic rings and hydrophobic alkyl chains. The amphiphilic nature of the phenolic lipids contributes to the formation of stable monomolecular layers in vitro (Kozubek and Tyman, 1999). Phenolic lipids also exhibit antimicrobial and antioxidation activities. In this review article, type III PKSs producing phenolic lipids in two microorganisms, Azotobacter vinelandii and S. griseus, are mainly described.

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The ars Operon in Azotobacter vinelandii

A. vinelandii is a Gram-negative, nitrogen-fixing soil bacterium that differentiates into metabolically dormant cysts under adverse environmental conditions (Lin and Sadoff, 1968). Cyst central body is surrounded by a thick membrane composed of a modified capsule (intine) and a layered outer shell (exine) (> Fig. 1a). Phenolic lipids such as alkylresorcinols and alkylpyrones (> Fig. 1b) are the major lipids in the cyst membrane. The major compound of the phenolic lipids is 5-henicosylresorcinol, which has a saturated C21 alkyl side chain (Reusch and Sadoff, 1983). The phenolic lipids in A. vinelandii presumably allow the cysts to resist desiccation. The genome database predicted the presence of an operon, named ars (alkylresorcinol synthesis) operon. This operon consists of two type III PKSs, arsB and arsC, and two type I fatty acid synthases (FASs), arsA and arsD (> Fig. 1c). In vitro study showed that ArsB and ArsC are an alkylresorcinol synthase and an alkylpyrone synthase, respectively (Funa et al., 2006). ArsB catalyzes three condensation of malonyl-CoA with an acyl starter substrate and subsequently cyclizes the resulting tetraketide intermediate via aldol condensation to yield an alkylresorcinol. On the other hand, ArsC catalyzes two or three condensation of malonyl-CoA with an acyl starter substrate and subsequently cyclizes the resulting triketide or tetraketide intermediate via lactonization to alkylpyrones. Moreover ArsB and ArsC use acyl substrates synthesized by ArsA and ArsD to produce phenolic lipids (Miyanaga et al., 2008). The lack of phenolic lipid synthesis caused by disruption of the ars operon showed that the ars operon is essential for the biosynthesis of phenolic lipids. The ars disruptant showed the formation of severely impaired cysts, indicating that the phenolic lipids are essential for formation of the cyst membrane (> Fig. 1a).

. Figure 1 Phenolic lipids produced by type III PKSs in A. vinelandii. (a) Electron micrographs of an ultrathin section of the cyst of A. vinelandii OP wild-type cell and an ars mutant cell. (b) Structures of alkylresorcinols and alkylpyrones produced in A. vinelandii. (c) Gene organization of the ars operon.

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The srs Operon in Streptomyces griseus

The genome sequence of S. griseus (Ohnishi et al., 2008) led to the discovery of an operon containing another type III PKS, in addition of RppA, named an srs operon (Streptomyces resorcinol synthesis). This operon consists of three genes: srsA encoding a type III PKS, srsB encoding a methyltransferase, and srsC encoding a flavoprotein hydroxylase (> Fig. 2a). Overexpression of part and all of the srs genes in a heterologous host, Streptomyces lividans, showed that SrsA is responsible for the synthesis of alkylresorcinols, SrsB catalyzes the methylation of alkylresorcinols to yield alkylresorcinol methyl ethers, and SrsC catalyzes the hydroxylation of the alkylresorcinol methyl ethers to yield alkylquinones (> Fig. 2b) (Funabashi et al., 2008). In vitro analysis showed that SrsA condenses one methylmalonyl-CoA and two malonyl-CoAs with an acyl substrate and subsequently cyclizes the resulting tetraketide

. Figure 2 Phenolic lipids produced by a type III PKS in S. griseus. (a) Gene organization of the srs operon in S. griseus and distribution of an srs-like operon in a wide variety of bacteria. (b) Structures of alkylresorcinols produced in S. griseus.

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intermediate via aldol condensation to yield an alkylresorcinol. A unique catalytic property of SrsA is that it synthesizes alkylresorcinols with an aromatic ring having a C-methyl group that is derived from methylmalonyl-CoA. Because the alkylresocinol methyl ethers and alkylquinones are detected mainly in the cell wall/membrane fraction of S. griseus, these phenolic lipids are associated with the cytoplasmic membrane. An srs disruptant, which produces no phenolic lipids, is highly sensitive to b-lactam antibiotics, such as penicillin G and cephalexin, inhibitors of the peptidoglycan synthesis, suggesting that the phenolic lipids confer penicillin resistance on S. griseus. The phenolic lipids may affect the characteristics and rigidity of the cell wall/cytoplasmic membrane. The wide distribution of a pair of genes encoding SrsA- and SrsB-like proteins among Gram-positive and negative bacteria (> Fig. 2a) prompts us to speculate that the phenolic lipids play a significant role in the biological membrane in various bacteria. For example, a purple photosynthetic bacterium Rhodospirillum centenum differentiates into cysts, like A. vinelandii (Berleman and Bauer, 2004). R. centenum contains a single type III PKS (chsA) gene accompanied by a methyltransferase gene, like the srs operon. Together with the fact that chsA gene expression substantially increases as R. centenum cells enter the phase of cyst cell development (Berleman et al., 2004), this operon is perhaps involved in the biosynthesis of phenolic lipids in the cyst membrane, as is for A. vinelandii.

4

Other Type III PKSs Responsible for Phenolic Lipid Synthesis

In vitro study by Saxena et al. (2003) showed that two type III PKS proteins, PKS11 and PKS18, in Mycobacterium tuberculosis prefer long acyl chain substrates and have the ability to produce alkylpyrones. However, no metabolites that are assumed to be derived from pyronecontaining long-chain compounds have been isolated from M. tuberculosis. Therefore, their biological function has not been elucidated. Filamentous fungi also contain type III PKS genes. A type III PKS from Neurospora crassa, ORAS (2’-oxoalkylresorcinolic acid synthase), was characterized (Funa et al., 2007). In vitro study showed that ORAS has the ability to produce alkylresorcylic acids from long acyl substrates. However, ORAS deletions gave no apparent phenotypic changes. Moreover, the production of alkylresorcinolic acids and alkylresorcinols could not be detected in N. crassa, probably because of a small amount of these polyketides produced. In a plant Sorghum bicolor, a PKS, probably belonging to the type III PKS family, was shown to be responsible for the biosynthesis of sorgoleone, which is an alkylquinone and is responsible for the allelopathic property (Dayan et al., 2003). Recently, the presence of a type III PKS was predicted by an expressed sequence tag (EST) analysis (Baerson et al., 2008). A methyltransferase (SbOMT3) and a hydroxylase are also involved in sorgoleone biosynthesis, as observed for the phenolic lipid synthesis in S. griseus. Phenolic lipids found in bacteria, fungi and plants are thus biosynthesized by type III PKSs. The biological functions of the alkylresorcinols as essential membrane lipids for encystment in A. vinelandii (Funa et al., 2006) and as probable membrane lipids conferring penicillin resistance on S. griseus (Funabashi et al., 2008) suggest that these phenolic lipids play an important role as minor components in the biological membrane in various bacteria. In fact, a computer search in the databases predicts the existence of srs- and ars-like operons in a wide variety of Gram-positive and negative bacteria. Among the srs-like operons in bacteria, we have determined the catalytic properties of the operon in Bacillus subtilis, which directs the synthesis of pyrone-type lipids (manuscript in preparation). srs-like genes are present not only

523

524

39

Type III Polyketide Synthases Responsible for Phenolic Lipid Synthesis

in prokaryotes but also in eukaryotes, including ORAS in the filamentous fungi and at least three such genes in the rice plant Oryza sativa (our unpublished results). Alkylresorcinols and alkylpyrones are therefore important, but thus far overlooked, membrane lipids in bacteria, fungi, and plants.

5

Research Needs

Many type III PKSs have been predicted by genome mining. Some products synthesized by type III PKSs exhibit various important biological activities. Type III PKSs are also appreciated for their natural diversity and architectural simplicity. Recent studies have shown that some type III PKSs play an important role in the biosynthesis of the biological membrane components in bacteria. Further studies on other unexploited type III PKSs will shed more light on novel biological components and bioactive substances.

References Austin MB, Noel JP (2003) The chalcone synthase superfamily of type III polyketide synthases. Nat Prod Rep 20: 79–110. Austin MB, Saito T, Bowman ME, Haydock S, Kato A, Moore BS, Kay RR, Noel JP (2006) Biosynthesis of Dictyostelium discoideum differentiation-inducing factor by a hybrid type I fatty acid-type III polyketide synthase. Nat Chem Biol 2: 494–502. Baerson SR, Dayan FE, Rimando AM, Nanayakkara NP, Liu CJ, Schro¨der J, Fishbein M, Pan Z, Kagan IA, Pratt LH, Cordonnier-Pratt MM, Duke SO (2008) A functional genomics investigation of allelochemical biosynthesis in Sorghum bicolor root hairs. J Biol Chem 283: 3231–3247. Bangera MG, Thomashow LS (1999) Identification and characterization of a gene cluster for synthesis of the polyketide antibiotic 2,4-diacetylphloroglucinol from Pseudomonas fluorescens Q2–87. J Bacteriol 181: 3155–63. Berleman JE, Bauer CE (2004) Characterization of cyst cell formation in the purple photosynthetic bacterium Rhodospirillum centenum. Microbiology 150: 383–390. Berleman JE, Hasselbring BM, Bauer CE (2004) Hypercyst mutants in Rhodospirillum centenum identify regulatory loci involved in cyst cell differentiation. J Bacteriol 186: 5834–5841. Dayan FE, Kagan IA, Rimando AM (2003) Elucidation of the biosynthetic pathway of the allelochemical sorgoleone using retrobiosynthetic NMR analysis. J Biol Chem 278: 28607–28611. Funa N, Awakawa T, Horinouchi S (2007) Pentaketide resorcylic acid synthesis by type III polyketide

synthase from Neurospora crassa. J Biol Chem 282: 14476–14481. Funa N, Ohnishi Y, Fujii I, Shibuya M, Ebizuka Y, Horinouchi S (1999) A new pathway for polyketide synthesis in microorganisms. Nature 400: 897–899. Funa N, Ozawa H, Hirata A, Horinouchi S (2006) Phenolic lipid synthesis by type III polyketide synthases is essential for cyst formation in Azotobacter vinelandii. Proc Natl Acad Sci 103: 6356–6361. Funabashi M, Funa N, Horinouchi S (2008) Phenolic lipids synthesized by type III polyketide synthase confer penicillin resistance on Streptomyces griseus. J Biol Chem 283: 13983–13991. Lin LP, Sadoff HL (1968) Encystment and polymer production by Azotobacter vinelandii in the presence of b-hydroxybutyrate. J Bacteriol 95: 2336–2343. Kozubek A, Tyman JH (1999) Resorcinolic lipids, the natural non-isoprenoid phenolic amphiphiles and their biological activity. Chem Rev 99: 1–26. Miyanaga A, Funa N, Awakawa T, Horinouchi S (2008) Direct transfer of starter substrates from type I fatty acid synthase to type III polyketide synthases in phenolic lipid synthesis. Proc Natl Acad Sci 105: 871–876. Ohnishi Y, Ishikawa J, Hara H, Suzuki H, Ikenoya M, Ikeda H, Yamashita A, Hattori M, Horinouchi S (2008) Genome sequence of the streptomycinproducing microorganism Streptomyces griseus IFO 13350. J Bacteriol 190: 4050–4060. Pfeifer V, Nicholson GJ, Ries J, Recktenwald J, Schefer AB, Shawky RM, Schro¨der J, Wohlleben W, Pelzer S (2001) A polyketide synthase in glycopeptide biosynthesis: the biosynthesis of the non-proteinogenic

Type III Polyketide Synthases Responsible for Phenolic Lipid Synthesis amino acid (S)-3,5-dihydroxyphenylglycine. J Biol Chem 276: 38370–38377. Reusch RN, Sadoff HL (1983) Novel lipid components of the Azotobacter vinelandii cyst membrane. Nature 302: 268–270.

39

Saxena P, Yadav G, Mohanty D, Gokhale RS (2003) A new family of type III polyketide synthases in Mycobacterium tuberculosis. J Biol Chem 278: 44780–44790.

525

40 Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria R. Kalscheuer Howard Hughes Medical Institute and Albert Einstein College of Medicine, Price Center, Bronx, NY, USA [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 528

2

The Key Enzyme of Bacterial Wax Ester and Triacylglycerol Biosynthesis: WS/DGAT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 528

3 3.1 3.2 3.3

Genes Involved in Precursor Formation for WE and TAG Biosynthesis . . . . . . . . . . 533 Biosynthesis of Diacylglycerols (DAGs) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 533 Biosynthesis of Fatty Alcohols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 533 Formation of Acyl-Coenzyme A Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 534

4

Alternative TAG Biosynthetic Routes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 534

5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 534

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_40, # Springer-Verlag Berlin Heidelberg, 2010

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40

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

Abstract: Triacylglycerols (TAGs) and wax esters (WEs) are abundant storage lipids in certain groups of eubacteria. The biochemical reactions involved in TAG and WE biosynthesis are very similar to those described for eukaryotes. However, the genes and encoded enzymes involved are fundamentally different between pro- and eukaryotes. This chapter summarizes the current knowledge of the genetic basis of TAG and WE biosynthesis in bacteria including a novel class of acyltransferase and pathways for precursor formation.

1

Introduction

Triacylglycerols (TAGs) are the predominant storage lipids present in eukaryotic microorganisms as well as in higher eukaryotes. Bacteria, however, were thought in the past to use only specialized polymeric neutral lipids, the polyhydroxyalkanoic acids, as their typical intracellular lipophilic reserve compounds. Sporadic descriptions of the occurrence of low- molecular weight storage lipids like wax esters (WEs) and TAGs in prokaryotes were regarded as rare exotic exemptions. The last decade, however, has dramatically changed our view of WE and TAG in bacteria and has revealed tremendous knowledge of the biochemical and genetic basis of biosynthesis of these compounds. These efforts were driven by interest in potential biotechnological applications but also by the intriguing possibility that these storage lipids might play an important role in virulence of some pathogenic bacteria, namely Mycobacterium tuberculosis and other mycobacteria.

2

The Key Enzyme of Bacterial Wax Ester and Triacylglycerol Biosynthesis: WS/DGAT

The final step in both TAG and WE biosynthesis in bacteria is mediated by a recently discovered novel acyltransferase class which is phylogenetically not related to those involved in formation of similar lipids in eukaryotes (Wa¨ltermann et al., 2007). The first gene of this enzyme class was identified in Acinetobacter baylyi ADP1 (Kalscheuer and Steinbu¨chel, 2003). The atfA gene codes for a highly unspecific acyltransferase that exhibits both WE synthase and acyl-coenzyme A:diacylglycerol acyltransferase activity (WS/DGAT). It requires fatty acid thioesters of coenzyme A (acyl-CoA) as activated substrates and can utilize diacylglycerols (DAGs) as well as fatty alcohols as acceptor molecules, resulting in both TAG and WE formation at the same time (> Fig. 1). The key role of WS/DGAT for storage lipid accumulation in A. baylyi ADP1 is demonstrated by the complete lack of WEs and drastic reduction in TAG levels in an atfA knock-out mutant. The extraordinary broad substrate range of AtfA has sparked research harnessing this enzyme for synthesis of a large variety of fatty acid esters for biotechnological applications (Sto¨veken and Steinbu¨chel, 2008), including a synthetic pathway for fuel production in recombinant bacteria (Kalscheuer et al., 2006). The atfA gene in A. baylyi ADP1 is not clustered with genes of any known importance for lipid metabolism (> Fig. 2). WS/DGAT homologues are present in genome sequences of numerous Gram-negative and -positive bacteria as well as in one archaea. They are particularly abundant in certain bacterial groups, namely actinomycetes and marine g-proteobacteria (> Table 1) which correlates well with frequent reports on presence of TAGs and/or WEs in members of these

. Figure 1 Pathways for biosynthesis of TAGs (a) and WEs (b + c) in A. baylyi ADP1 and other oleogenous bacteria. G3P, glycerol-3-phosphate.

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

40 529

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40

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

. Figure 2 Organization of WS/DGAT gene loci with proven pivotal importance for WE and/or TAG biosynthesis. WS/DAGT genes are shown in black.

groups (Alvarez and Steinbu¨chel, 2002; Kalscheuer et al., 2007). In addition to A. baylyi, the pivotal role of WS/DGAT enzymes for TAG and/or WE biosynthesis has been demonstrated for several other bacteria by means of gene deletion mutants supported by in vitro enzyme activity data: Mycobacterium tuberculosis (Sirakova et al., 2006), Alcanivorax borkumensis (Kalscheuer et al., 2007) and Streptomyces coelicolor (Arabolaza et al., 2008). Many bacteria harbor multiple WS/DGAT genes which is especially articulate in mycobacteria with up to 16 homologues (> Table 1). Although several of these genes have been shown to encode active acyltransferases, they are not fully functionally redundant, and a single WS/DGAT enzyme seems to be the clearly predominant contributor to WE and/or TAG accumulation in all cases analyzed so far (tgs1 in M. tuberculosis, atfA1 in A. borkumensis, sco0958 in S. coelicolor ; > Fig. 2) (Sirakova et al., 2006; Kalscheuer et al., 2007; Arabolaza et al., 2008). The role of those WS/DGAT genes that are apparently not required for lipid accumulation under the tested conditions is unresolved. They may be important for lipid accumulation under yet unknown conditions, or they could be involved in biosynthesis of some unidentified lipid molecules. The extensive group of WS/DGAT genes in M. tuberculosis suggests an important function of these acyltransferases for virulence of this pathogen. In fact, prominent lipid inclusions consisting of TAGs have been observed in M. tuberculosis cells in sputum from patients with active disease, and stored TAGs are believed to contribute to long-term survival in the host (Garton et al., 2008). Tgs1 is the main DGAT enzyme for TAG biosynthesis in M. tuberculosis under various in vitro conditions as revealed by mutational analysis (Sirakova et al., 2006). The tgs1 gene is a member of the DosR regulon which is responsive to stresses resembling those supposedly present in vivo in the human host like hypoxia, low pH and nitrosative stress. Exposure to such stresses results in strong induction of tgs1 which is paralleled by increased accumulation of TAGs (Sirakova et al., 2006). However, there are no reports available so far whether loss of TAG biosynthesis as a result of tgs1 deletion has any influence on virulence or long-term persistence of M. tuberculosis in animal models.

40

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

. Table 1 Occurrence of WS/DGAT genes in bacterial genomes

Group Archaea

Strain

Number of homologues

Natronomonas pharaonis DSM 2160

1

Sphingopyxis alaskensis RB2256

2

Gram-negative a-Proteobacteria

b-Proteobacteria

g-Proteobacteria

d-Proteobacteria

Bradyrhizobium japonicum USDA 110

1

Erythrobacterr sp. NAP1

1

Erythrobacter litoralis HTCC2594

1

Erythrobacter sp. SD-21

1

Limnobacter sp. MED105

2

Rhodoferax ferrireducens T118

3

Polaromonas sp. JS666

2

Methylibium petroleiphilum PM1

1

Polaromonas naphthalenivorans CJ2

1

Acinetobacter baylyi ADP1

1

Acinetobacter baumannii ATCC 17918

1

Psychrobacter arcticus 273-4

1

Psychrobacter cryohalolentis K5

1

Psychrobacter sp. PRwf-1

2

Hahella chejuensis KCTC 2396

4

Alkanivorax borkumensis SK2

2

Marinobacter aquaeolei VT8

5

Marinobacter algicola DG893

4

Marinobacter sp. ELB17

1

Congregibacter litoralis KT71

3

Gamma-proteobacterium HTCC2080

1

Moritella sp. PE36

4

Photobacterium profundum SS9

1

Photobacterium profundum 3TCK

1

Alteromonas macleodii

1

Aeromonas hydrophila subsp. hydrophila ATCC 7966

1

Aeromonas salmonicida subsp. salmonicida A449

1

Marine gamma-proteobacterium HTCC2143

1

Reinekea sp. MED297

1

Stigmatella aurantiaca DW4/3-1

1

Myxococcus xanthus DK 1622

1

Plesiocystis pacifica SIR-1

1

Anaeromyxobacter sp. Fw109-5

1

531

532

40

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

. Table 1 (Continued) Group

Number of homologues

Strain Anaeromyxobacter dehalogenans 2CP-C

1

Gram-positive Actinobacteria

Acidothermus cellulolyticus 11B

1

Janibacter sp. HTCC2649

1

Frankia sp. EAN1pec

1

Mycobacterium avium 104

10

Mycobacterium avium subsp. paratuberculosis K-10 Mycobacterium bovis AF2122/97

12

Mycobacterium bovis BCG str. Pasteur 1173P2

13

Mycobacterium gilvum PYR-GCK

10

Mycobacterium leprae TN

1

Mycobacterium smegmatis mc 155 2

Bacteroidetes/ Chlorobi

Other

8

6

Mycobacterium sp. JLS

16

Mycobacterium sp. KMS

15

Mycobacterium sp. MCS

15

Mycobacterium tuberculosis C

12

Mycobacterium tuberculosis CDC1551

13

Mycobacterium tuberculosis F11

13

Mycobacterium tuberculosis H37Ra

15

Mycobacterium tuberculosis H37Rv

13

Mycobacterium ulcerans Agy99

12

Mycobacterium vanbaalenii PYR-1

13

Nocardia farcinia IFM 10152

5

Nocardioides sp. JS614

5

Rhodococcus sp. RHA1

13

Saccharopolyspora erythraea NRRL 2338

2

Streptomyces coelicolor A3(2)

3

Streptomyces avermitilis MA-4680

1

Microscilla marina ATCC 23134

1

Polaribacter dokdonensis MED152

1

Salinibacter ruber DSM 13855

1

Solibacter usitatus Ellin6076

1

Roseiflexus sp. RS-1

2

Roseiflexus castenholzii DSM 13941

2

Acidobacteria bacterium Ellin345

1

Genes were identified using AtfA from A. baylyi ADP1 as reference by BLASTP search within 874 finished microbial genome sequences available online at http://www.ncbi.nlm.nih.gov

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

40

3

Genes Involved in Precursor Formation for WE and TAG Biosynthesis

3.1

Biosynthesis of Diacylglycerols (DAGs)

Formation of DAGs used as substrate for TAG biosynthesis occurs by reactions that are partially shared with glycerophospholipid biosynthesis (> Fig. 1a, the so-called Kennedy pathway). In the first step, glycerol-3-phosphate (G3P) is acylated to 1-acyl-G3P by the G3P acyltransferase PlsB which can utilize fatty acid thioesters of either acyl carrier protein (ACP) derived from fatty acid de novo biosynthesis (acyl-ACP) or acyl-CoA as substrates. However, many bacteria are lacking a plsB gene. Here the first acylation step is mediated by a recently discovered alternative pathway (Zhang and Rock, 2008). PlsX is a transacylase that forms acylphosphate from acyl-ACP which is then used as activated substrate by the acyl-phosphate:G3P acyltransferase PlsY. While the PlsX-PlsY pathway is present in some Gram-negative storage lipid accumulating bacteria, most TAG and WE synthesizing species, particularly actinomycetes, seem to rely on the PlsB reaction since plsX and plsY genes are lacking. Regardless of the pathway, the formed 1-acyl-G3P is then acylated by the 1-acyl-G3P acyltransferase PlsC which is present in virtually all bacteria leading to synthesis of phosphatidic acid. Phosphatidic acid is the metabolic branch point dividing TAG and phospholipid biosynthesis in bacteria since it can be converted to CDP-diacylglycerol which serves as precursor for the formation of membrane glycerophospholipids (Zhang and Rock, 2008). Finally, DAG is released from phosphatidate by dephosphorylation mediated by phosphatidate phosphatase (PAP). This is the first reaction of the Kennedy pathway that is specific to TAG biosynthesis in bacteria. PAP genes have been identified in various eukaryotes (Carman and Han, 2006) but they do not possess homologues in bacterial genomes. Thus, a bacterial PAP gene still awaits identification.

3.2

Biosynthesis of Fatty Alcohols

The long-chain fatty alcohols required for WE biosynthesis are formed from the corresponding acyl-CoAs by two consecutive reduction steps (> Fig. 1b). Acyl-CoA is reduced by the acyl-CoA reductase Acr1 to the corresponding fatty aldehyde. The acr1 gene was first identified in A. baylyi ADP1 by analysis of transposon-induced WE deficient mutants, and it codes for a NADPH-dependent enzyme (Reiser and Somerville, 1997). Like the atfA gene in this bacterium, the acr1 gene is not clustered with genes of any known relevance for lipid metabolism. In the second step, the fatty aldehydes are further reduced to fatty alcohols by a NADPH-dependent fatty aldehyde reductase. However, a gene coding for this reductase could not be identified so far. Additionally, fatty alcohols are generated as an intermediate during aerobic catabolism of long-chain n-alkanes (> Fig. 1c, for details refer to > Chapter 3, Vol. 2, Part 1), and WE accumulation in alkane-degrading bacteria like Acinetobacter and Alkanivorax is well documented (Ishige et al., 2003).

533

534

40 3.3

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

Formation of Acyl-Coenzyme A Substrates

In addition to DAG or fatty alcohols as acceptor molecules, the WS/DGAT reaction requires acyl-CoA as an activated substrate. Acyl-CoAs are derived from exogenous fatty acids or oxidation of aliphatic hydrocarbons via conversion by acyl-CoA synthetase FadD. Since WS/DGAT enzymes are apparently unable to efficiently utilize acyl-ACP, these end products of fatty acid de novo biosynthesis need to be converted to the corresponding CoA thioesters. This can be achieved through hydrolysis of acyl-ACP by thioesterases releasing free fatty acids which are subsequently converted to acyl-CoA by FadD. Alternatively, this could theoretically also be mediated by a direct acyl-ACP:CoA transacylation. However, such a transacylase activity has never been described in bacteria. Regardless of the pathway, a key feature of lipid accumulating bacteria is their ability to deregulate fatty acid de novo biosynthesis and to provide a high intracellular acyl-CoA pool. Fatty acid biosynthesis is strictly regulated and tightly coupled to membrane lipid biosynthesis in non-lipid accumulating bacteria (Zhang and Rock, 2008). The regulatory mechanisms and pathways by which oleogenous prokaryotes uncouple fatty acid biosynthesis from phospholipid biosynthesis and achieve high-level acylCoA formation are not understood.

4

Alternative TAG Biosynthetic Routes

Mutational studies as described above indicate that acyltransferases of the WS/DGAT family are the key enzymes in storage lipid biosynthesis probably in all WE and TAG accumulating bacteria and account for the by far major part of neutral lipids. However, residual levels of TAGs as well as low but significant DGAT activity still present in these WS/DGAT mutants clearly indicate that alternative, non-WS/DGAT-type acyltransferases must be present which might contribute to TAG formation to a minor extent. This phenotype has been observed in the atfA mutant of A. baylyi ADP1 (Kalscheuer and Steinbu¨chel, 2003), the atfA1 atfA2 double mutant of A. borkumensis SK2 (Kalscheuer et al., 2007) and the sco0958 sco1280 sco0123 triple mutant of S. coelicolor (Arabolaza et al., 2008). Those DGAT isoenzymes have not been identified yet. DGAT isoenzymes are known in eukaryotes, but no homologues are present in bacterial genomes. In S. coelicolor, not only DGAT isoenzmyes but an alternative acyl-CoA independent TAG biosynthesis route might be present. Phospholipid:diacylglycerol acyltransferase (PDAT) mediates acyl-CoA-independent TAG production in yeast and plants, and recently PDAT activity has been also detected in S. coelicolor (Arabolaza et al., 2008). However, there are no homologues of eukaryotic PDATs in the genome of S. coelicolor or any other bacterium. Thus, a bacterial PDAT gene remains to be identified and the contribution of this alternative pathway to total TAG accumulation is unknown.

5

Research Needs

Although the key step in bacterial TAG and WE biosynthesis mediated by WS/DGAT has been identified, genes of essential steps in precursor formation such as PAP and fatty aldehyde reductase are still unknown. Understanding how fatty acid biosynthesis is deregulated in oleogenous bacteria will be critical in establishment vital processes for biotechnological

Genetics of Wax Ester and Triacylglycerol Biosynthesis in Bacteria

40

production of designer lipids using recombinant strains. Of medical relevance, it will be important to determine whether TAG metabolism is critical for virulence and persistence of M. tuberculosis and if this metabolic process offers opportunities for developing new approaches of antituberculous chemotherapy.

References Alvarez HM, Steinbu¨chel A (2002) Triacylglycerols in prokaryotic microorganisms. Appl Microbiol Biotechnol 60: 367–376. Arabolaza A, Rodriguez E, Altabe S, Alvarez H, Gramajo H (2008) Multiple pathways for triacylglycerol biosynthesis in Streptomyces coelicolor. Appl Environ Microbiol 74: 2573–2582. Carman GM, Han GS (2006) Roles of phosphatidate phosphatase enzymes in lipid metabolism. Trends Biochem Sci 31: 694–699. Garton NJ, Waddell SJ, Sherratt AL, Lee SM, Smith RJ, Senner C, Hinds J, Rajakumar K, Adegbola RA, Besra GS, Butcher PD, Barer MR (2008) Cytological and transcript analyses reveal fat and lazy persisterlike bacilli in tuberculous sputum. PLoS Med 5: e75. Ishige T, Tani A, Sakai Y, Kato N (2003) Wax ester production by bacteria. Curr Opin Microbiol 6: 244–250. Kalscheuer R, Steinbu¨chel A (2003) A novel bifunctional wax ester synthase/acyl-CoA:diacylglycerol acyltransferase mediates wax ester and triacylglycerol biosynthesis in Acinetobacter calcoaceticus ADP1. J Biol Chem 278: 8075–8082. Kalscheuer R, Sto¨lting T, Steinbu¨chel A (2006) Microdiesel: Escherichia coli engineered for fuel production. Microbiology 152: 2529–2536. Kalscheuer R, Sto¨veken T, Malkus U, Reichelt R, Golyshin PN, Sabirova JS, Ferrer M, Timmis KN,

Steinbu¨chel A (2007) Analysis of storage lipid accumulation in Alcanivorax borkumensis: Evidence for alternative triacylglycerol biosynthesis routes in bacteria. J Bacteriol 189: 918–928. Reiser S, Somerville C (1997) Isolation of mutants of Acinetobacter calcoaceticus deficient in wax ester synthesis and complementation of one mutation with a gene encoding a fatty acyl coenzyme A reductase. J Bacteriol 170: 2969–2975. Sirakova TD, Dubey VS, Deb C, Daniel J, Korotkova TA, Abomoelak B, Kolattukudy PE (2006) Identification of a diacylglycerol acyltransferase gene involved in accumulation of triacylglycerol in Mycobacterium tuberculosis under stress. Microbiology 152: 2717–2725. Sto¨veken T, Steinbu¨chel A (2008) Bacterial acyltransferases as an alternative for lipase-catalyzed acylation for the production of oleochemicals and fuels. Angew Chem Int Ed Engl 47: 3688–3694. Wa¨ltermann M, Sto¨veken T, Steinbu¨chel A (2007) Key enzymes for biosynthesis of neutral lipid storage compounds in prokaryotes: properties, function and occurrence of wax ester synthases/ acyl-CoA: diacylglycerol acyltransferases. Biochimie 89: 230–242. Zhang YM, Rock CO (2008) Membrane lipid homeostasis in bacteria. Nat Rev Microbiol 6: 222–233.

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41 Players in the Neutral Lipid Game – Proteins Involved in Neutral Lipid Metabolism in Yeast K. Athenstaedt Institute of Biochemistry, University of Technology Graz, Graz, Austria [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 538

2 Enzymes Catalyzing Neutral Lipid Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 539 2.1 Triacylglycerol Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 539 2.2 Steryl Ester Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 540 3

Mutations Leading to Structural Changes in Lipid Particles . . . . . . . . . . . . . . . . . . . . . 540

4 Neutral Lipid Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 541 4.1 Triacylglycerol Lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 542 4.2 Degradation of Steryl Esters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 542 5

Key Knowledge Gaps and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 544

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41

Players in the Neutral Lipid Game – Proteins Involved in Neutral Lipid Metabolism in Yeast

Abstract: Synthesis, storage and degradation of neutral lipids enable cells to continue cell metabolism when nutrients are no longer provided by the environment. Major neutral lipids occurring in yeast are triacylglycerols and steryl esters. These hydrophobic molecules are sequestered from the cytosolic environment in the core of so-called lipid particles (lipid droplets). When nutrients are no longer provided by the environment, hydrolytic enzymes catalyze the degradation of triacylglycerols and steryl esters. The respective breakdown products serve as energy source and/or building blocks for membrane formation. Here, enzymes catalyzing neutral lipid synthesis and degradation in the budding yeast Saccharomyces cerevisiae are described with special emphasis to their localization and regulation. Formation of lipid particles is not only disturbed in cells defective in polypeptides directly involved in neutral lipid synthesis and degradation but also in a number of mutants defective in pathways which are not obviously linked to neutral lipid turnover. Although research over the past decade provided major insights into neutral lipid metabolism, many aspects of neutral lipid synthesis, storage and degradation remain to be elucidated. Research needs for a better understanding of neutral lipid turnover are outlined at the end of this chapter.

1

Introduction

A detailed knowledge about enzymes catalyzing the formation and degradation of neutral lipids, the regulation of these enzymes as well as the correlation of different pathways with neutral lipid metabolism is required for biomedical purposes as well as studies in the field of biotechnology. Besides a general understanding of neutral lipid metabolism, investigations in neutral lipid turnover are aimed at finding targets for altering the accumulation of storage molecules. In humans excessive accumulation of neutral lipids is related to several severe diseases such as arteriosclerosis, obesity and diabetes type II, thus scientists are seeking for means how to avoid detrimental accumulation of storage molecules. In contrast, genetic manipulations resulting in high amounts of triacylglycerols (TAG) are in focus of biotechnological research, since in industry microorganisms such as the oleaginous yeast Yarrowia lipolytica are used for, e.g., single cell oil production and production of nutrients enriched in essential fatty acids. In the budding yeast Saccharomyces cerevisiae as well as in other eukaryotes TAG and steryl esters (SE) constitute the major portion of neutral lipids. Investigations with the model organism yeast identified numerous polypeptides involved in neutral lipid metabolism. For many of these proteins counterparts in other microorganisms as well as in higher eukaryotes have been identified, thus indicating that lipid metabolism is well conserved across the different kingdoms of life. In Saccharomyces cerevisiae four enzymes catalyze the formation of SE and TAG. Since substantial amounts of neutral lipids cannot be incorporated into the phospholipid bilayer, these hydrophobic molecules are sequestered in so-called lipid particles, cell compartments destined for storage of these compounds. The formation of lipid particles is not only disturbed in mutants defective in enzymes catalyzing neutral lipid synthesis and degradation but also in strains blocked in other pathways which are not obviously connected to lipid metabolism. In yeast six hydrolytic enzymes have been identified catalyzing the degradation of TAG and SE. The breakdown products of neutral lipid degradation are sterols, diacylglycerol and fatty acids which serve as building blocks for membrane formation and/or for energy production. For general information about neutral lipid synthesis, storage and degradation the reader is

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41

referred to > Chapter 35 of this volume and some recent reviews covering specifically neutral lipid metabolism in yeast (Czabany et al., 2007; Rajakumari et al., 2008).

2

Enzymes Catalyzing Neutral Lipid Synthesis

2.1

Triacylglycerol Synthesis

Yeast synthesizes TAG via an acyl-CoA dependent as well as an acyl-CoA independent mechanism (> Chapter 35, Vol. 1, Part 5). The first TAG synthase identified in Saccharomyces cerevisiae has been Lro1p, catalyzing TAG synthesis independent of acyl-CoA (Dahlqvist et al., 2000; Oelkers et al., 2000). Lro1p has been identified due to its homology to the mammalian lecithin:cholesterol acyltransferase (LCAT). Like its mammalian homolog, Lro1p uses glycerophospholipids as acyl-donors but transfers the acyl chain to the sn-3 position of diacylglycerol instead. In vitro Lro1p exhibits a preference for unsaturated fatty acids bound to the sn-2 position of phosphatidylethanolamine and phosphatidylcholine (Ghosal et al., 2007). Enzymatic measurements revealed the presence of Lro1p in the endoplasmic reticulum (ER) (Dahlqvist et al., 2000; Oelkers et al., 2000). A transmembrane domain predicted at the N-terminus of Lro1p is most probably responsible for the localization of Lro1p to the ER, since a truncated version of Lro1p lacking the first 98 amino acids (including the transmembrane domain) fused to alpha-factor secretion signal is efficiently secreted into the medium upon heterologous expression in Pichia pastoris (Ghosal et al., 2007). The truncated Lro1-protein is highly active indicating that the N-terminal transmembrane domain is only required for membrane localization but not for enzymatic activity. Lro1p is a glycosylated protein and deglycosylation significantly reduces its enzymatic activity (Ghosal et al., 2007). Due to the glycosylation of Lro1p it is assumed that the bulk of the polypeptide faces the luminal side of the ER. Lro1p is the only acyl-CoA independent TAG synthase of Saccharomyces cerevisiae, since in a strain lacking LRO1 TAG are exclusively formed via the acyl-CoA dependent pathway. The major acyl-CoA dependent TAG synthase of Saccharomyces cerevisiae is Dga1p. Enzymatic measurements revealed a 70- to 90-fold enrichment of acyl-CoA dependent TAG synthase activity in lipid particles over homogenate which is ascribed to Dga1p (Sorger and Daum, 2002). However, Dga1p is also present in the ER (Oelkers et al., 2002; Sorger and Daum, 2002). Similar to Lro1p, a transmembrane domain is predicted at the N-terminus of Dga1p. However, whereas the active site of Lro1p is assumed to be oriented to the luminal side of the ER, that of Dga1p is most likely exposed to the cytosol (Sta˚hl et al., 2004). These differences in orientation may lead to different TAG pools being physically separated and having different physiological functions. Moreover, Lro1p and Dga1p are active at different growth stages of the cell. Whereas Lro1p is the main contributor to TAG synthesis in cells of the exponential growth phase, the latter enzyme is highly active in cells of the stationary phase (Oelkers et al., 2002). Overexpression of DGA1 and enhanced leucine biosynthesis in a snf2D mutant background but not in wild-type significantly increases TAG accumulation (Kamisaka et al., 2007). Since the protein levels of Dga1p are similar in both respective mutants, it is hypothesized that post-translational modifications of Dga1p account for the higher amount of TAG in the snf2D mutant overexpressing DGA1. By contrast, overexpression of LRO1 in snf2D cells reduces TAG accumulation. Snf2p is a transcription factor forming a part of the SWI/SNF (switching/

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sucrose non-fermenting) chromatin remodeling complex. Since SNF2 disruption markedly decreases the expression of genes encoding the phospholipid-biosynthetic enzymes regulated by inositol and choline (Kodaki et al., 1995), a defect in Snf2p may significantly reduce the amount of glycerophospholipids which serve as a substrate for Lro1p. Alternatively, Lro1p may compensate the defect in phospholipid synthesis by catalyzing the reverse reaction, namely transferring an acyl-chain from TAG to a lyso-phospholipid. In wild-type cells, Lro1p and Dga1p together account for 90% of TAG synthase activity (Oelkers et al., 2002; Sandager et al., 2002). Are1p and Are2p, two acyl-CoA:cholesterol acyltransferase (ACAT) related enzymes (see below), have been shown to be responsible for the residual TAG synthase activity in lro1Ddga1D cells. Like Dga1p, both Are1p and Are2p are catalyzing TAG synthesis via the acyl-CoA dependent mechanism. However, the preferred substrate of Are1p and Are2p are sterols, thus these two enzymes are referred as acyl-CoA dependent SE synthases.

2.2

Steryl Ester Synthesis

In contrast to higher eukaryotic organisms, Saccharomyces cerevisiae catalyzes the esterification of sterols only in presence of acyl-CoA. In the budding yeast Are1p and Are2p, two acyl-CoA:cholesterol acyltransferase (ACAT) related enzymes, catalyze the formation of SE (Yang et al., 1996; Yu et al., 1996). Since SE cannot be detected in a mutant deleted of both ARE1 and ARE2, the respective gene products are the only SE synthases of yeast. Both Are1p and Are2p are localized to the ER (Zweytick et al., 2000). Different substrate specificities may account for the redundancy of SE synthesizing enzymes in the ER. Whereas Are1p has a significant preference for sterol precursors, ergosterol is the main substrate of Are2p. A defect in ARE2 is reflected by the reduction of the total amount of SE to 26% of control (Yang et al., 1996; Yu et al., 1996). In contrast, in an are1D mutant the cellular SE level remains nearly unaffected. This is not surprising having in mind that ergosterol is the most abundant sterol in wild-type cells. The higher contribution of Are2p to SE formation is paralleled by a significantly higher transcription activity through the ARE2 promoter as well as a 12 times higher half-life of the respective mRNA compared to ARE1 (Jensen-Pergakes et al., 2001). Under anaerobic conditions, however, the major portion of SE is formed by Are1p. This can be explained by the fact that ergosterol biosynthesis is an oxygen consuming process and limitation of oxygen leads to the accumulation of sterol precursors which are the preferred substrate of Are1p.

3

Mutations Leading to Structural Changes in Lipid Particles

As outlined above, in the budding yeast Saccharomyces cerevisiae four enzymes, namely Lro1p, Dga1p, Are1p and Are2p, contribute to the formation of TAG and SE. Neutral lipids synthesized by these polypeptides are sequestered from the cytosolic environment forming the hydrophobic core of so-called lipid particles (> Chapter 35, Vol. 1, Part 5). A quadruple mutant deleted of LRO1, DGA1, ARE1 and ARE2 is viable but lacks lipid particles. By contrast, triple mutant cells expressing only one of these four genes contain lipid particles revealing that the activity of a single neutral lipid synthesizing enzyme is sufficient for lipid particle

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formation. All triple mutants contain a high portion of lipid particles with a diameter between 0.3 and 0.4 mm which is similar to wild-type particles (Czabany et al., 2008). However, whereas in wild-type cells the size distribution of lipid particles is rather narrow, it is significantly altered in cells harboring only one active neutral lipid synthesizing enzyme. As an example, in lro1Dare1Dare2D cells (i.e., Dga1p is active) 30% of the lipid particles range in diameter from 0.45 to 0.55 mm. Since in addition this mutant contains an equal number of smaller lipid particles with a diameter between 0.25 and 0.35 mm, the mean diameter of the particles is similar to wild-type. Cells of all other triple mutants contain a significant higher portion of smaller lipid particles ranging in diameter from 0.25 to 0.35 mm. The appearance of lipid particles is also altered in a mutant defective in the major TAG lipase Tgl3p (see below). In a tgl3D mutant the total amount of TAG is threefold increased compared to control (Athenstaedt and Daum, 2003). Staining cells of this mutant with the lipophilic fluorescent dye Nile Red specific for lipid particles leads to increased lipid particle fluorescence (Fei et al., 2008). However, not only defects in processes obviously linked to neutral lipid turnover cause alterations in lipid particle formation, but also defects in several other metabolic pathways. As an example, most recently Henry and coworkers (Gaspar et al., 2008) provided evidence that a block in the secretory pathway leads to lipid particle proliferation caused by increased TAG formation. The link between the secretory pathway and TAG synthesis is phosphatidic acid (PtdOH), the key intermediate for the formation of all glycerolipids. Dephosphorylation of PtdOH leads to diacylglycerol, the direct precursor of TAG synthesis. Activation of PtdOH by CTP, on the other hand, yields CDP-diacylglycerol the precursor for the formation of all glycerophospholipids. In cells defective in secretion membrane material (glycerophospholipids) in form of vesicles cannot exit the ER and may thus cause secretory stress to the ER. By favoring dephosphorylation of PtdOH over activation with CTP further conversion of PtdOH is rerouted from glycerophospholipid synthesis towards TAG formation, thus providing a degree of protection to cells during secretory stress. Several mutants containing lipid particles with altered appearance have been obtained by a screen aimed to identify genes which are required for sterol homeostasis (Fei et al., 2008). Detailed investigations revealed for some of these mutants a link between neutral lipid synthesis and secretory glycosylation. As an example, in cells defective in CAX4 and SEC53, respectively, the number of lipid particles is significantly reduced. Both Cax4p and Sec53p are involved in N-linked glycosylation. By contrast, a higher number of lipid particles has been observed in ume6D and cdc50D cells. The gene product of UME6 is a transcriptional regulator inducing early meiotic genes and regulating specific mitotic genes. Cdc50p has been shown to genetically interact with sterol biosynthetic genes (Kishimoto et al., 2005). However, this protein is also involved in cell polarization and regulates the cellular localization of Drs2p, a lipid translocase (Natarajan et al., 2004; Saito et al., 2004). These few examples already demonstrate that lipid particle formation is influenced by a number of processes which are not obviously linked to neutral lipid turnover.

4

Neutral Lipid Degradation

Degradation of TAG and SE is catalyzed by hydrolytic enzymes, namely TAG lipases and SE hydrolases. The breakdown products formed by these catabolic processes are sterols,

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diacylglycerol and free fatty acids. Whereas sterols can be directly integrated into membrane bilayers, the latter two molecules serve as building blocks for membrane lipid formation. Alternatively, b-oxidation of fatty acids supplies the cell with energy.

4.1

Triacylglycerol Lipases

In Saccharomyces cerevisiae TAG degradation is accomplished by three TAG lipases, namely Tgl3p, Tgl4p and Tgl5p (Athenstaedt and Daum, 2003; 2005). All three TAG lipases are located to lipid particles, thus close to their substrate. Whereas in comparison to wild-type cells a defect in TGL3 is reflected by a threefold increased TAG level, TAG accumulation is only moderately increased in tgl4D cells and not changed at all in tgl5D cells. This can be explained by different substrate preferences of Tgl3p, Tgl4p and Tgl5p. Tgl3p is a rather unspecific TAG lipase regarding the fatty acids cleaved off of TAG. Tgl4p, on the other hand prefers myristic (C14:0) and palmitic (C16:0) acid, and Tgl5p exhibits a preference for hexacosanoic (C26:0) acid. However, all three TAG lipases are highly specific for TAG and do not hydrolyze SE. Tgl3p has been the first TAG lipase identified of a lipase protein family which contains polypeptides with the GXSXG motif characteristic for lipolytic enzymes but lacking additional homologies to other previously known lipases. By now this protein family not only comprises Tgl3p, Tgl4p and Tgl5p of yeast but also TAG lipases of mammals, insects and plants (reviewed in Athenstaedt and Daum, 2006). To date, information about regulatory aspects of TAG degradation is rather limited. However, a mechanism for regulating the activity of TAG lipases seems to be likely, since otherwise unspecific degradation of TAG may occur. Preliminary results revealed that Tgl4p is a substrate of cyclin-dependent kinase Cdk1p, thus linking lipolysis to cell cycle progression (Kohlwein and Petschnigg, 2007). Most interestingly, the phosphatidate phosphatase Pah1p which catalyzes the degradation of PtdOH yielding diacylglycerol the direct precursor of TAG, is also controlled by Cdk1p, however, in the opposite way (O’Hara et al., 2006). Whereas phosphorylation of Tgl4p through Cdk1p activates the lipase, phosphorylation of Pah1p renders this protein inactive resulting in a higher amount of PtdOH which is channeled towards glycerophospholipid synthesis. Thus, simultaneous phosphorylation of Tgl4p and Pah1p contributes to cell proliferation.

4.2

Degradation of Steryl Esters

Similar to TAG degradation, three enzymes catalyze the deacylation of SE, namely Tgl1p, Yeh1p and Yeh2p (Jandrositz et al., 2005; Ko¨ffel et al., 2005; Mu¨llner et al., 2005). Whereas Tgl1p and Yeh1p are located to lipid particles in yeast, Yeh2p has been detected in the plasma membrane. Analysis of SE in a yeh2D mutant revealed a slightly higher level of zymosterol, thus indicating a preference of Yeh2p for this sterol precursor (Mu¨llner et al., 2005). In yeast, anaerobic conditions can be mimicked by heme deficiency. Investigations of mutants deleted of two out of the three genes encoding Tgl1p, Yeh1p and Yeh2p, and the triple deletion mutant tgl1Dyeh1Dyeh2D in a heme deficient background indicated that Yeh1p is the major enzyme catalyzing SE degradation under anaerobic conditions (Ko¨ffel and Schneiter, 2006) (> Table 1).

SE hydrolase

SE hydrolase

SE hydrolase

Tgl1p

Yeh1p

Yeh2p

SE

SE

SE (TAG)

TAG

TAG

TAG

Acyl-CoA, sterols (diacylglycerol)

Acyl-CoA, sterols (diacylglycerol)

Acyl-CoA, diacylglycerol

Phospholipids, diacylglycerol

Substrate

ER endoplasmic reticulum; LP lipid particle; PM plasma membrane; SE steryl ester; TAG triacylglycerol

TAG lipase

Tgl5p

Acyl-CoA:sterol acyltransferase

Are2p

TAG lipase

Acyl-CoA:sterol acyltransferase

Are1p

Tgl4p

Acyl-CoA:diacylglycerol acyltransferase

Dga1p

TAG lipase

Phospholipid:diacylglycerol acyltransferase

Main molecular function

Lro1p

Gene product

Degradation Tgl3p

Synthesis

Involved in

Induced upon anaerobic conditions

Glycosylation

Regulation/ Modification

Sterol, fatty acid

Sterol, fatty acid

Sterol, fatty acid

Diacylglycerol, fatty acid Induced upon anaerobic conditions

Diacylglycerol, fatty Phosphorylation acid

Diacylglycerol, fatty acid

SE (TAG)

SE (TAG)

TAG

TAG

Product

. Table 1 Polypeptides involved in neutral lipid metabolism in the budding yeast Saccharomyces cerevisiae

PM

LP

LP

LP

LP

LP

ER

ER

ER, LP

ER

Localization

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Players in the Neutral Lipid Game – Proteins Involved in Neutral Lipid Metabolism in Yeast

Key Knowledge Gaps and Research Needs

Although to date major players in neutral lipid turnover in yeast are identified, the answers to several questions related to neutral lipid synthesis, storage and degradation are still missing. As outlined above, enzymes catalyzing neutral lipid synthesis and degradation occur in redundancy. This leads to the question why at least two enzymes exist which are catalyzing essentially the same reaction. One simple explanation is that isoenzymes allow ongoing turnover even if one of these polypeptides is defective. However, this seems to be only half of the truth. It is hypothesized that each isoenzyme has also a specific function which makes it singular. As an example, whereas the TAG mobilization defect of tgl3D cells can at least to some extent be compensated by the additional TAG lipases Tgl4p and Tgl5p, the homologs of Tgl3p are unable to reverse the sporulation defect of a homozygous diploid tgl3D/tgl3D mutant (Athenstaedt and Daum, 2003). This demonstrates the indispensable role of Tgl3p in spore formation. For several isoenzymes involved in neutral lipid turnover a unique function remains to be elucidated. This will not only significantly contribute to our current understanding of neutral lipid metabolism, but furthermore reveal some ‘‘novel’’ relationships of neutral lipid synthesis, storage and degradation with other cell metabolic functions. One approach which has a high potential to answer the question concerning the existence of enzymes with overlapping function is studying regulatory aspects of neutral lipid metabolism. Furthermore, by investigating regulation at the gene level as well as the protein level some unexpected links between neutral lipid turnover and other cell metabolic processes may be indicated. As an example, a link between neutral lipid degradation, glycerophospholipid synthesis and cell cycle progression has been observed by studying the regulation of the TAG lipase Tgl4p (see above). Since currently information about regulatory aspects is rather limited, one major aim of future investigations has to be the elucidation of these aspects. As outlined in detail above, all enzymes catalyzing neutral lipid synthesis are localized to the ER. The major acyl-CoA dependent TAG synthase Dga1p, however, is also present on lipid particles. The dual localization of Dga1p leads to several questions. One question concerns the mechanism by which Dga1p reaches the lipid particle surface. Is it by a budding process as suggested by the budding model for lipid particle biogenesis (> Chapter 35, Vol. 1, Part 5)? Why is it Dga1p, which is dually localized, and not one of the other neutral lipid synthesizing enzymes? In vitro measurements revealed that Dga1p is active in both ER and lipid particles. However, to date we can only speculate whether this is also true in vivo. An active Dga1p on lipid particles may be required to compensate random degradation of TAG which may occur since all TAG lipases are equally localized to the particle. In a mutant deleted of the major TAG lipase Tgl3p the amount of TAG is significantly increased suggesting that Dga1p localized to lipid particles is indeed active in vivo. This observation leads to several additional questions, e.g., is TAG formed by Dga1p the preferred substrate of Tgl3p? How is Tgl3p prevented from hydrolyzing just synthesized TAG? Or is there a concomitant synthesis and degradation of TAG? What would be the sense of such a permanent turnover? Already those few questions/hypotheses mentioned above reveal that many aspects of neutral lipid metabolism are currently far from being understood and require further investigation. However, with our detailed knowledge about the features of single polypeptides governing neutral lipid metabolism we are already well equipped to set the next steps: to elucidate the interplay of enzymes and organelles during neutral lipid turnover and to unravel the significance of neutral lipid synthesis, storage and degradation for other cell metabolic processes.

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significantly increase lipid accumulation in the Dsnf2 disruptant of Saccharomyces cerevisiae. Biochem J 408: 61–68. Kishimoto T, Yamamoto T, Tanaka K (2005) Defects in structural integrity of ergosterol and the Cdc50pDrs2p putative phospholipid translocase cause accumulation of endocytic membranes, onto which actin patches are assembled in yeast. Mol Biol Cell 16: 5592–5609. Kodaki T, Hosaka K, Nikiwa J, Yamashita S (1995) The SNF2/SWI2/GAM1/TYE3/RIC1 gene is involved in the coordinate regulation of phospholipid synthesis in Saccharomyces cerevisiae. J Biochem (Tokyo) 117: 362–368. Ko¨ffel R, Schneiter R (2006) Yeh1 constitutes the major steryl ester hydrolase under heme-deficient conditions in Saccharomyces cerevisiae. Eukaryot Cell 5: 1018–1025. Ko¨ffel R, Tiwari R, Falquet L, Schneiter R (2005) The Saccharomyces cerevisiae YLL012/YEH1, YLR020/ YEH2, and TGL1 genes encode a novel family of membrane-anchored lipases that are required for steryl ester hydrolysis. Mol Cell Biol 25: 1655–1668. Kohlwein SD, Petschnigg P (2007) Lipid-induced cell dysfunction and cell death: lessons from yeast. Curr Hypertens Rep 9: 455–461. Mu¨llner H, Deutsch G, Leitner E, Ingolic E, Daum G (2005) YEH2/YLR020c encodes a novel steryl ester hydrolase of the yeast Saccharomyces cerevisiae. J Biol Chem 280: 13321–13328. Natarajan P, Wang J, Hua Z, Graham TR (2004). Drs2pcoupled aminophospholipid translocase activity in yeast Golgi membranes and relationship to in vivo function. Proc Natl Acad Sci USA 101: 10614–10619. Oelkers P, Cromley D, Padamsee M, Billheimer JT, Sturley SL (2002) The DGA1 gene determines a second triglyceride synthetic pathway in yeast. J Biol Chem 277: 8877–8881. Oelkers P, Tinkelenberg A, Erdeniz N, Cromley D, Billheimer J, Sturley SL (2000) A lecithin cholesterol acyltransferase-like gene mediates diacylglycerol esterification in yeast. J Biol Chem 275: 15609–15612. O’Hara L, Han GS, Peak-Chew S, Grimsey N, Carman GM, Siniossoglou S (2006) Control of phospholipid synthesis by phosphorylation of the yeast lipin Pah1p/Smp2p Mg2+-dependent phosphatidate phosphatase. J Biol Chem 281: 34537–34548. Rajakumari S, Grillitsch K, Daum G (2008) Synthesis and turnover of non-polar lipids in yeast. Prog Lipid Res 47: 157–171. Saito K, Fujimura-Kamada K, Furuta N, Kato U, Umeda M, Tanaka K (2004) Cdc50p, a protein

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required for polarized growth, associates with the Drs2p P-type ATPase implicated in phospholipid translocation in Saccharomyces cerevisiae. Mol Biol Cell 15: 3418–3432. Sandager L, Gustavsson MH, Sta˚hl U, Dahlqvist A, Wiberg E, Banas´ A, Lenman M, Ronne H, Stymne S (2002) Storage lipid synthesis is nonessential in yeast. J Biol Chem 277: 6478–6482. Sorger D, Daum G (2002) Synthesis of triacylglycerols by the acyl-coenzyme A:diacyl-glycerol acyltransferase Dga1p in lipid particles of the yeast Saccharomyces cerevisiae. J Biol Chem 184: 519–524. Sta˚hl U, Carlsson AS, Lenman M, Dahlqvist A, Wiberg E, Banas´ W, Banas´ A, Stymne S (2004) Cloning and characterization of a phospholipid:diacylglycerol

acyltransferase from Arabidopsis. Plant Physiol 135: 1324–1335. Yang H, Bard M, Bruner DA, Gleeson A, Deckelbaum RJ, Aljinovic G, Pohl TM, Rothstein R, Sturley SL (1996) Sterol esterification in yeast: a two-gene process. Science 272: 1353–1356. Yu C, Kennedy NJ, Chang CCY, Rothblatt J (1996) Molecular cloning and characterization of two isoforms of Saccharomyces cerevisiae acyl-CoA:sterol acyltransferases. J Biol Chem 271: 24157–24163. Zweytick D, Leitner E, Kohlwein SD, Yu C, Rothblatt J, Daum G (2000) Contribution of Are1p and Are2p to steryl ester synthesis in the yeast Saccharomyces cerevisiae. Eur J Biochem 267: 1075–1082.

Part 7

The Microbes Section Editor: Terry McGenity

42 Taxonomy of Methanogens Y. Liu Department of Microbiology, University of Georgia, Athens, GA, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 550 2 Taxonomy of Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 550 3 Phylogeny of Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 551 4 Ecology of Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 556 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 556

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_42, # Springer-Verlag Berlin Heidelberg, 2010

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42

Taxonomy of Methanogens

Abstract: Methanogens are strictly anaerobic, methane-producing Archaea. They all belong to the phylum Euryarchaeota. Although methanogens share a set of physiological characteristics, they are phylogenetically very diverse. The current taxonomy classifies methanogens into five well-established orders: Methanobacteriales, Methanococcales, Methanomicrobiales, Methanosarcinales, and Methanopyrales. This taxonomy is supported by 16S rRNA gene sequences as well as a number of physiological properties, e.g., substrates for methanogenesis, nutritional requirements, morphologies, and structures of cell envelopes. The 16S rRNA gene sequence analysis of the strain SANAE, a representative of the clone lineage Rice Cluster I, suggests that it represents a novel order of methanogens, Methanocellales. Methanogens are abundant in a wide variety of anaerobic environments where they catalyze the terminal step in the anaerobic food chain by converting methanogenic substrates into methane. The complexity of methanogenesis pathways suggests an ancient monophyletic origin of methanogens, a hypothesis, which is supported by phylogenetic analyses based upon DNA sequences.

1

Introduction

Methanogens are microorganisms that produce methane as the end-product of their anaerobic respiration. All methanogens share three common features. (1) They are obligate methane producers, obtaining all or most of their energy for growth from producing large quantities of methane. (2) They are archaea, belonging to the phylum Euryarchaeota. (3) They are strict anaerobes, limiting their growth to anaerobic environments. Methanogens can only utilize a restricted number of substrates for methane production or methanogenesis. The substrates are limited to three major types: CO2 + H2 or a few other electron donors such as formate, methyl-group-containing compounds, and acetate. Methanogens using these three types of substrates are classified as hydrogenotrophs, methylotrophs, and acetotrophs, respectively. Most organic substances, for instance, carbohydrates, proteins, and long-chain fatty acids and alcohols, are not substrates for methanogenesis. Exceptions are that some hydrogenotrophs can also use secondary alcohols, such as 2-propanol, 2-butanol, and cyclopentanol, as electron donors. A small number can use ethanol (Bleicher et al., 1989; Frimmer and Widdel, 1989; Widdel, 1986; Widdel et al., 1988). However, even these organic compounds, which can obviously be assimilated, are only incompletely oxidized to ketones (secondary alcohols) and acetate (ethanol), and methane is derived from CO2 reduction. Methanogenesis is a complex process that requires a number of unique enzyme complexes and unusual coenzymes (reviewed by Hedderich and Whitman, 2006). Although the methanogenesis pathways of the three nutritional groups start differently, the final steps leading to methane are common in virtually all methanogens. The bioenergetics of methanogenesis employs both proton and sodium gradients generated by primary pumps for ATP synthesis. Due to the complexity of methanogenesis, all common methanogens are expected to originate from an ancient ancestor.

2

Taxonomy of Methanogens

Although methanogens are united by a few common features, they are phylogenetically diverse. The taxonomy of methanogens that has been developed in the last 3 decades has aimed to reflect the phylogenetic diversity of methanogens and be consistent with the taxonomy

Taxonomy of Methanogens

42

of other prokaryotes (Balch et al., 1979; Boone et al., 1993; Whitman et al., 2001). An overview of the current taxonomy of methanogens is given in > Table 1. Organisms from different orders have less than 82% 16S rRNA sequence similarity. Organisms with less than 88–93% and less than 93–95% 16S rRNA sequence similarity are separated into different families and genera, respectively. Organisms are distinguished as separate species if their DNA reassociation is less than 70%, the change in the melting temperature of their hybrid DNA is greater than 5
C, and substantial phenotypic differences exist (Stackebrandt et al., 2002; Wayne et al., 1987). When 16S rRNA data are available, organisms with a similarity of less than 98% are considered as separate species. However, sequence similarity of greater than 98% is not considered as sufficient evidence that two organisms belong to the same species. Methanogens are currently classified into five orders: Methanobacteriales, Methanococcales, Methanomicrobiales, Methanosarcinales, and Methanopyrales (Whitman et al., 2001, 2006). This taxonomy is supported by comparative 16S rRNA gene sequence analyses as well as distinctive phenotypic properties, such as different cell envelope structure, lipid composition, and substrate range. Some representative characteristics are listed in > Table 2 and further described in following chapters. A novel order of methanogen, Methanocellales, was proposed recently (Sakai et al., 2008). This order is currently represented by a single strain, Methanocella paludicola strain SANAE, which was originally isolated from rice paddy soil (Sakai et al., 2007). Based on comparative 16S rRNA gene sequence analysis, strain SANAE is affiliated with Rice Cluster I (RC-I), a clone lineage within the radiation of Methanosarcinales and Methanomicrobiales (review in Conrad et al., 2006). The 16S rRNA gene sequence similarities between strain SANAE and members of Methanosarcinales range from 80% to 82.8%, and those between the strain and members of Methanomicrobiales range from 77.5% to 82.4% (Sakai et al., 2008). This novel strain utilizes H2 + CO2 or formate as methanogenesis substrates. The cells are nonmotile and rod shaped. The growth temperature ranges from 25
C to 40
C, with an optimum of 35–37
C. The pH for growth ranges from 6.5 to 7.8, with an optimum of 7.0. The salinity for growth ranges from 0% to 0.1% (w/v) NaCl. It is physiologically distinguished from members of Methanosarcinales, which can use acetate and methylated compounds for methanogenesis. However, the phenotypic distinction between strain SANAE and members of Methanomicrobiales needs further investigation.

3

Phylogeny of Methanogens

All modern methanogens share the same set of homologous enzymes and cofactors required for methanogenesis, suggesting an ancient monophyletic origin of methanogens. In the phylogenetic tree based on 16S rRNA gene sequences, methanogens are separated into two major groups (> Fig. 1). The Class I methanogens include Methanobacteriales, Methanococcales, and Methanopyrales, and the Class II methanogens include Methanomicrobiales and Methanosarcinales (Bapteste et al., 2005). Non-methanogenic lineages, Archaeoglobales, Halobacteriales, and Thermoplasmatales, are interspersed in the tree. Three hypotheses are proposed to explain this branching of methanogens. 1. Methanogens and these non-methanogen lineages shared a common ancestor, and genes required for methanogenesis were lost in these non-methanogens. This hypothesis is supported by the presence of a few genes encoding methanogenesis enzymes in the genome of Archaeoglobus fulgidus but is challenged by aerobic growth in both the Halobacteriales

551

M. cuniculi, M. stadtmanae M. defluvii, M. marburgensis, M. thermoautotrophicus, M. thermoflexus, M. thermophilus, M. wolfeii

Methanosphaera

Methanothermobacter

M. okinawensis, M. thermolithotrophicus

Methanothermococcus

Methanospirillaceae

M. cariaci, M. frigidum, M. marinum, M. organophilum M. paynteri M. mobile M. endosymbiosus, M. limicola, M. petrolearius

Methanogenium

Methanolacinia

Methanomicrobium

Methanoplanus

M. hungatei

M. aquaemaris, M. formosanus, M. liminatans, M. tationis

Methanofollis

Methanospirillum

M. bourgensis, M. chikugoensis, M. marisnigri, M. palmolei, M. submarinus, M. thermophilus

M. formicicus, M. igneus

Methanoculleus

Methanotorris

M. fervens, M. indicus, M. infernus, M. jannaschii, M. vulcanius

M. aeolicus, M. maripaludis, M. vannielii, M. voltae

Methanococcus

M. fervidus, M. sociabilis

M. acididurans, M. arboriphilus, M. curvatus, M. cuticularis, M. filiformis, M. gottschalkii, M. millerae, M. olleyae, M. oralis, M. ruminantium, M. smithii, M. thaueri, M. woesei, M. wolinii

Methanobrevibacter

Methanothermus

M. aarhusense, M. alcaliphilum, M. beijingense, M. bryantii, M. congolense, M. espanolae, M. formicicum, M. ivanovii, M. oryzae, M. palustre, M. subterraneum, M. uliginosum

Speciesa

Methanobacterium

Genus

Methanocaldococcaceae Methanocaldococcus

Methanococcaceae

Methanothermaceae

Methanobacteriaceae

Methanomicrobiales Methanomicrobiaceae

Methanococcales

Methanobacteriales

Family

42

Order

. Table 1 Taxonomy of methanogens

552 Taxonomy of Methanogens

b

M. halophilus, M. mahii, M. portucalensis M. bombayensis, M. oregonensis, M. taylorii, M. tindarius, M. vulcani

Methanohalophilus

Methanolobus

Methanopyraceae

Methanocellaceae

Methanocellales

Methanocella

Methanopyrus

Methanosaeta

M. paludicola

M. kandleri

M. shengliensis

M. concilii, M. harundinacea, M. thermophila

M. zhilinae

Methanosalsum

Methermicoccus

M. blatticola

Methanimicrococcusb

Methermicoccaceae

Type species of the genera are in bold Placement in higher taxon is tentative

a

M. evestigatum

Methanohalobium

Methanomethylovorans M. hollandica, M. thermophila

M. alaskense, M. burtonii, M. methylutens

Methanococcoides

M. palustris

Candidatus Methanosphaerula

M. acetivorans, M. baltica, M. barkeri, M. lacustris, M. mazei, M. semesiae, M. siciliae, M. thermophila, M. vacuolata

M. boonei

Candidatus Methanoregula

Methanosarcina

M. tarda

Methanolinea

Methanosaetaceae

Methanosarcinaceae

Methanopyrales

Methanosarcinales

Unclassified

M. chunghsingensis, M. halotolerans, M. pumilus, M. taiwanensis

M. bavaricum, M. labreanum, M. parvum, M. sinense

Methanocalculusb

Methanocorpusculaceae Methanocorpusculum

Taxonomy of Methanogens

42 553

Pseudomurein nd

nd, not determined a Major substrates utilized for methanogenesis. Parentheses means utilized sometimes b Compounds can be contained in cellular lipids, depending on the species c Except the genus Methanothermus

H2 + CO2, formate

+

Rods

Methanocellales

Protein, glycoprotein

Protein, glycoprotein

Protein

Pseudomurein, protein

Cell wall



Rods

Methanopyrales

H2 + CO2

Methanol, methylamine,  acetate (H2 + CO2)

Pseudosarcina, cocci, sheathed rods

+/

+

Methanosarcinales

H2 + CO2, formate



H2 + CO2, formate (secondary alcohols)

Cocci

Methanococcales

H2 + CO2 (formate, CO, methanol, secondary alcohols)

c

Motility

Methanomicrobiales Cocci, rods, spirals, sheathed rods

Rods, cocci

Shape

Methanobacteriales

Order

Glucose, galactose, aminopentanetetrol, glycerol

Glucose, N-acetylglucosamine, serine, ethanolamine

Glucose, N-acetylglucosamine, myo-inositol, ethanolamine, serine

Polar lipids

nd

Archaeol

nd

nd

Archaeol, hydroxyarchaeol, Glucose, galactose, mannose, caldarchaeol myo-inositol, ethanolamine, serine, glycerol

Archaeol, caldarchaeol

Archaeol, caldarchaeol, hydroxyarchaeol, macrocyclic archaeol

Caldarchaeol, archaeol, hydroxyarchaeol

Core lipids

Cellular lipidsb

42

Methanogenesis substratesa

. Table 2 Some characteristics of the methanogen orders

554 Taxonomy of Methanogens

Taxonomy of Methanogens

42

. Figure 1 Phylogenetic tree for the methanogenic archaea and other euryarchaeotes based upon 16S rRNA sequences. The alignment was manually edited to include 1,251 positions. The tree was constructed with the neighbor-joining algorithm in MEGA4. Bootstrap analysis was performed with 1,000 replicates, and values greater than 50% are labeled on the nodes. The scale bar is 0.05 expected nucleotide substitutions per site. The 16S rRNA sequence of Methanosalsum zhilinae was according to Mathrani et al. (1988). The GenBank accession numbers for the other sequences are given following their names.

555

556

42

Taxonomy of Methanogens

and Thermoplasmatales. This hypothesis also suggests that the common ancestor of Euryarchaeota was a methanogen (Gribaldo and Brochier-Armanet, 2006). 2. Methanogenesis in various branches was acquired by horizontal gene transfer (HGT). However, the core genes required for methanogenesis are not linked on the genomes of methanogens, thus the simultaneous acquisition via lateral transfer is unlikely, and the transfer of single genes would not confer a selective advantage (Gribaldo and BrochierArmanet, 2006). 3. The phylogeny based on 16S rRNA gene is misleading, and methanogens and Archaeoglobus shared a common ancestor exclusive of all other archaea. This hypothesis is supported by a recent phylogenomics analysis showing that ten proteins are exclusively shared in methanogens and A. fulgidus (Gao and Gupta, 2007), while no proteins are exclusively shared in methanogens and any of the Halobacteriales or Thermoplasmatales (Gao and Gupta, 2007). Therefore, methanogens and Archaeoglobus appear to have a closer relationship within the Euryarchaeota, but this leaves open the question of the apparently paradoxical phylogenetic position of both the Halobacteriales and Thermplasmatales. Therefore, more genomic sequences of archaea are needed to prove this hypothesis.

4

Ecology of Methanogens

Methanogens are abundant in a wide variety of anaerobic habitats such as marine sediments, freshwater sediments, flooded soils, human and animal gastrointestinal tracts, anaerobic digestors, landfills, and geothermal systems (Liu and Whitman, 2008). In some natural habitats, methanogens are also present in micro-oxic environments. For example, members of Methanobrevibacter have been isolated from large dental caries and subgingival plaque in the human mouth and gut periphery in termites. They are also somewhat oxygen tolerant, probably due to the presence of catalase activity and the protection given by O2-utilizing microbes (Brusa et al., 1987; Belay et al., 1988; Leadbetter and Breznak, 1996). RC-I methanogens are prevalent in rice rhizosphere, which is transiently oxic, and a reconstituted RC-I genome encodes a unique set of antioxidant enzymes, which may explain an aerotolerant life style (Erkel et al., 2006). 3þ 2 are In methanogenic habitats, electron acceptors such as O2 ; NO 3 ; Fe ; and SO4 limiting. When electron acceptors other than CO2 are present, methanogens are outcompeted by the bacteria that utilize them. This phenomenon occurs mainly because the reduction of these compounds is thermodynamically more favorable than CO2 reduction to methane. However, because CO2 is generated during fermentations, it is seldom limiting in anaerobic environments. Besides methanogens, homoacetogens are another group of anaerobes that can reduce CO2 for energy production. However, acetogenesis with H2 is thermodynamically less favorable than methanogenesis. Therefore, homoacetogens do not compete well with methanogens in many habitats. However, homoacetogens outcompete methanogens in some environments, such as the hindgut of certain termites and cockroaches. Possible explanations may be their metabolic versatility as well as lower sensitivity to O2.

5

Research Needs

Recent culture-independent studies have revealed the presence of novel phylogenetic groups of methanogens. Their isolation and characterization will shed new insight into these organisms.

Taxonomy of Methanogens

42

For instance, investigations of rumen methanogens have found a novel lineage containing at least two families. The 16S rRNA gene sequences of this group have similarities closest to, but less than 80%, with those of Methanosarcinales (Nicholson et al., 2007). The Rice Cluster I is abundant in rice paddy soils, but only one strain has been isolated so far. Discovery and isolation of new strains will certainly add to our knowledge of the diversity of methanogens. Methanogens have fewer easily determined physiological characteristics than most bacteria. Comparative 16S rRNA gene sequence analyses are indispensable for determination of taxonomic levels higher than species. However, it is frequently insufficient for taxonomy of methanogens at species and subspecies levels. For instance, some isolates of Methanobrevibacter have >98% 16S rRNA gene sequence similarities but exhibit less than 50% DNA relatedness, suggesting that they belong to different species (Lin and Miller, 1998; Keswani and Whitman, 2001). The discovery of novel molecular markers is desirable. The methylcoenzyme M reductase alpha-subunit (mcrA) gene has been applied as a phylogenetic marker for methanogens in addition to 16S rRNA genes (Springer et al., 1995) and as a target for the detection of methanogens in a wide range of environments (Ohkuma et al., 1995; Lueders et al., 2001; Luton et al., 2002; Earl et al., 2003; Kemnitz et al., 2004). Phylogenomic analyses based upon whole-genome sequences may lead to improvement of the taxonomy and better view of phylogenetic relationships. A more complete database of methanogen genome sequences is required for this purpose.

References Balch WE, Fox GE, Magrum LJ, Woese CR, Wolfe RS (1979) Methanogens: reevaluation of a unique biological group. Microbiol Mol Biol Rev 43: 260–296. Bapteste E, Brochier C, Boucher Y (2005) Higher-level classification of the Archaea: evolution of methanogenesis and methanogens. Archaea 1: 353–363. Belay N, Johnson R, Rajagopal BS, de Macario EC, Daniels L (1988) Methanogenic bacteria from human dental plaque. Appl Environ Microbiol 54: 600–603. Bleicher K, Zellner G, Winter J (1989) Growth of methanogens on cyclopentanol/CO2 and specificity of alcohol dehydrogenase. FEMS Microbiol Lett 59: 307–312. Boone DR, Whitman WB, Rouviere P (1993) Diversity and taxonomy of methanogens. In Methanogenesis: Ecology, Physiology, Biochemistry and Genetics. JG Ferry (ed.). New York: Chapman & Hall, pp. 35–80. Brusa T, Conca R, Ferrara A, Ferrari A, Pecchioni A (1987) The presence of methanobacteria in human subgingival plaque. J Clin Periodontol 14: 470–471. Conrad R, Erkel C, Liesack W (2006) Rice Cluster I methanogens, an important group of Archaea producing greenhouse gas in soil. Curr Opin Biotechnol 17: 262–267.

Earl J, Hall G, Pickup RW, Ritchie DA, Edwards C (2003) Analysis of methanogen diversity in a hypereutrophic lake using PCR-RFLP analysis of mcr sequences. Microb Ecol 46: 270–278. Erkel C, Kube M, Reinhardt R, Liesack W (2006) Genome of Rice Cluster I archaea – the key methane producers in the rice rhizosphere. Science 313: 370–372. Frimmer U, Widdel F (1989) Oxidation of ethanol by methanogenic bacteria. Arch Microbiol 152: 479–483. Gao B, Gupta R (2007) Phylogenomic analysis of proteins that are distinctive of Archaea and its main subgroups and the origin of methanogenesis. BMC Genomics 8: 86. Gribaldo S, Brochier-Armanet C (2006) The origin and evolution of Archaea: a state of the art. Phil Trans R Soc B 361: 1007–1022. Hedderich R, Whitman WB (2006) Physiology and biochemistry of the methane-producing Archaea. In The Prokaryotes, M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.). 3rd edn. New York: Springer-Verlag. Kemnitz D, Chin K-J, Bodelier P, Ralf Conrad (2004) Community analysis of methanogenic archaea within a riparian flooding gradient. Environ Microbiol 6: 449–461.

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42

Taxonomy of Methanogens

Keswani J, Whitman W (2001) Relationship of 16S rRNA sequence similarity to DNA hybridization in prokaryotes. Int J Syst Evol Microbiol 51: 667–678. Leadbetter JR, Breznak JA (1996) Physiological ecology of Methanobrevibacter cuticularis sp. nov. and Methanobrevibacter curvatus sp. nov., isolated from the hindgut of the termite Reticulitermes flavipes. Appl Environ Microbiol 62: 3620–3631. Lin C, Miller TL (1998) Phylogenetic analysis of Methanobrevibacter isolated from feces of humans and other animals. Arch Microbiol 169: 397–403. Liu Y, Whitman WB (2008) Metabolic, phylogenetic, and ecological diversity of the methanogenic archaea. Ann N Y Acad Sci 1125: 171–189. Lueders T, Chin K-J, Conrad R, Friedrich M (2001) Molecular analyses of methyl-coenzyme M reductase alpha-subunit (mcrA) genes in rice field soil and enrichment cultures reveal the methanogenic phenotype of a novel archaeal lineage. Environ Microbiol 3: 194–204. Luton PE, Wayne JM, Sharp RJ, Riley PW (2002) The mcrA gene as an alternative to 16S rRNA in the phylogenetic analysis of methanogen populations in landfill. Microbiology 148: 3521–3530. Mathrani IM, Boone DR, Mah RA, Fox GE, Lau PP (1988) Methanohalophilus zhilinae sp. nov., an alkaliphilic, halophilic, methylotrophic methanogen. Int J Syst Bacteriol 38: 139–142. Nicholson M, Evans P, Joblin K (2007) Analysis of methanogen diversity in the rumen using temporal temperature gradient gel electrophoresis: identification of uncultured methanogens. Microb Ecol 54: 141–150. Ohkuma M, Noda S, Horikoshi K, Kudo T (1995) Phylogeny of symbiotic methanogens in the gut of the termite Reticulitermes speratus. FEMS Microbiol Lett 134: 45–50. Sakai S, Imachi H, Sekiguchi Y, Ohashi A, Harada H, Kamagata Y (2007) Isolation of key methanogens for global methane emission from rice paddy fields: a novel isolate affiliated with the clone cluster Rice Cluster I. Appl Environ Microbiol 73: 4326–4331.

Sakai S, Imachi H, Hanada S, Ohashi A, Harada H, Kamagata Y (2008) Methanocella paludicola gen. nov., sp. nov., a methane-producing archaeon, the first isolate of the lineage ‘Rice Cluster I’, and proposal of the new archaeal order Methanocellales ord. nov. Int J Syst Evol Microbiol 58: 929–936. Springer E, Sachs MS, Woese CR, Boone DR (1995) Partial gene sequences for the A subunit of methylcoenzyme M reductase (mcrI) as a phylogenetic tool for the family Methanosarcinaceae. Int J Syst Bacteriol 45: 554–559. Stackebrandt E, Frederiksen W, Garrity GM, Grimont PAD, Kampfer P, Maiden MCJ, Nesme X, RosselloMora R, Swings J, Truper HG, Vauterin L, Ward AC, Whitman WB (2002) Report of the ad hoc committee for the re-evaluation of the species definition in bacteriology. Int J Syst Evol Microbiol 52: 1043–1047. Wayne LG, Brenner DJ, Colwell RR, Grimont PAD, Kandler O, Krichevsky MI, Moore LH, Moore WEC, Murray RGE (1987) Report of the ad hoc committee on reconciliation of approaches to bacterial systematics. Int J Syst Bacteriol 37: 463–464. Whitman W, Bowen T, Boone D (2006) The methanogenic bacteria. In The Prokaryotes, 3rd edn. M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.) New York: Springer-Verlag, pp. 165–207. Whitman WB, Boone DR, Koga Y, Keswani J (2001) Taxonomy of methanogenic archaea. In Bergey’s Mannual of Systematic Bacteriology, 2nd edn. DR Boone, RW Castenholtz, GM Garrity (eds.). New York: Springer-Verlag. 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, Rouvie`re PE, Wolfe RS (1988) Classification of secondary alcohol-utilizing methanogens including a new thermophilic isolate. Arch Microbiol 150: 477–481.

43 Methanobacteriales Y. Liu Department of Microbiology, University of Georgia, Athens, GA, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560 2 Descriptive Features . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560 3 Phylogeny and Taxonomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 560 4 Ecology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 569

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_43, # Springer-Verlag Berlin Heidelberg, 2010

560

43

Methanobacteriales

Abstract: The order Methanobacteriales is composed of two families, Methanobacteriaceae and Methanothermaceae, which are distinguished by 16S rRNA sequence similarities below 89% and differences in cell wall structure and growth temperatures. Members of the Methanobacteriaceae are thermophilic or mesophilic, and they possess pseudomurein as a major component of the cellular envelope. Members of the Methanothermaceae are hyperthermophilic and possess a protein surface layer in addition to the pseudomurein layer. Most members of this order produce methane by CO2 reduction with H2. Formate, CO, or secondary alcohols are used as alternative electron donors in some species.

1

Introduction

Members of the order Methanobacteriales use a limited range of substrates for methanogenesis. Most of them use H2 to reduce CO2 to CH4, except the genus Methanosphaera, which uses H2 to reduce methanol. Some members can also use formate, CO, or secondary alcohols as electron donors. Some species can grow autotrophically using CO2 as the sole carbon source, and some species are mixotrophs or heterotrophs, which may require acetate, amino acids, peptones, yeast extract, B-vitamins, and/or rumen fluid for growth. Ammonium is a major nitrogen source. Sulfide can serve as the sole sulfur source, and some species can reduce elemental sulfur to sulfide. Cells are generally rod-shaped with a length of 0.6–25 mm, often forming chains or filaments up to 40 mm in length. Cells typically stain Gram-positive, but the wall does not contain muramic acid. Pesudomurein is the predominant polymer in the cell wall. Members of the genus Methanothermus have double-layered cell wall, consisting of an inner pseudomurein layer and an outer S-layer composed of protein. The cellular lipids contain caldarchaeol, archaeol, and, in some species, hydoxyarchaeol as core lipids. The polar lipids can contain glucose, N-acetylglucosamine, myo-inositol, ethanolamine, and serine, depending on the species. Most species are nonmotile. However, members of the genus Methanothermus are motile via peritrichous flagella. The optimum growth temperatures of members of the Methanobacteriales vary from 20 to 88 C. The genus Methanothermus can grow at temperatures up to 97 C. The pH optima of members of Methanobacteriales vary from 5.5 to 9.

2

Descriptive Features

Descriptive properties of the Methanobacteriales are summarized in > Tables 1–5. Further information can be found in (Bonin and Boone, 2006; Boone et al., 2001).

3

Phylogeny and Taxonomy

Methanobacteriales are currently classified into two families and five genera based upon 16S rRNA sequences, DNA reassociation levels, and phenotypic characteristics. The family Methanobacteriaceae contains three mesophilic genera, Methanobacterium, Methanobrevibacter, and Methanosphaera, and one thermophilic genus Methanothermobacter. The family Methanothermaceae is represented by one hyperthermophilic genus, Methanothermus.

WeN4

8–2

M.o.H. Anaerobic digestor

C

GP9

MF

Ivanov Rock core

FPi

aarhusense

alcaliphilum

beijingense

bryantii

congolense

espanolae

formicicum

ivanovii

oryzae

Rice field

Sewage sludge

5–18

Cell length (mm)

0.3–0.4 3–10

0.5–0.8 1–15

0.4–0.8 2–15

3–22

0.4–0.5 2–10

0.5–1.0 10–15

0.4–0.5 3–5

0.5–0.6 2–25

0.7

Sludge 0.8 of a bleachcraft mill

Anaerobic digestor

Anaerobic digestor

Alkaline lake

Marine sediment

H2-LR

Species

Sourcea

Type strain

Cell width (mm)

H2 + CO2, formate

H2 + CO2

H2 + CO2, formate

H2 + CO2 (2-propanol, 2-butanol)

H2 + CO2 (2-propanol, 2-butanol, cyclopentanol)

H2 + CO2 (2-propanol, 2-butanol, cyclopentanol)

H2 + CO2, formate

H2 + CO2

H2 + CO2

Methanogenesis substratesb

None

None

None

ndd

None

None

YE

TP or YE

None

Required organic compounds

. Table 1 Descriptive characteristics of the species of the genus Methanobacterium

6.0–8.5 (7.0)

6.5–8.2 (7.0–7.4)

>10– < 55 (37–45)

4.6–7.0 (5.6–6.2)

5.9–8.2 (7.2)

6.6–7.8 (7–7.5)

20–42 (40)

0–3

nd

0.6–5.4

NaCl range (%, w/v)

0–2.5

0–1

nd

nd

nd

nd (6.9–7.2) nd

6.5–8.0 (7.2)

7.0–9.9 (8.1–9.1)

5–9 (7.5–8)

25–50 (37–45)

15–50 (35)

25–50 (37–42)

nd (37–39)

25–50 (37)

25–45 (37)

>5– < 48 (45)

Temperature range pH range (optimum) (optimum) (oC)

nd

16–18

13

10

7.5

nd

14

nd

nd

Doubling timec (h)

Reference

Worakit et al. (1986)

Boone (1987)

Boone (1987)

Patel et al. (1990)

31 (LC)

Joulian et al. (2000)

36.6 (Tm) Belyaev et al. (1983)

41–42 (Bd)

34 (Tm)

39.5 (LC) Cuzin et al. (2001)

33–38 (Bd)

38.9 (Tm) Ma et al. (2005)

57 (BD)

34.9 (LC) Shlimon et al. (2004)

GC content (mol%)

Methanobacteriales

43 561

F

A8p

P2St

Species

palustre

subterraneum

uliginosum

Marshy soil

Deep granitic ground water

Peat bog

Sourcea H2 + CO2, formate, 2-propanol (2-butanol)

Methanogenesis substratesb

H2 + CO2

0.6–1.2 H2 + CO2, formate

2.5–5

0.2–0.6 2–4

0.1– 0.15

0.5

Cell length (mm)

None

None

None

Required organic compounds

15–45 (37– 40)

3.6–45 (20–40)

20–45 (33– 37)

6.0–8.5 (6.0–7.5)

6.5–9.2 (7.8–8.8)

nd (7.0)

Temperature range pH range (optimum) (optimum) (oC)

nd

0–8

0–1.8

NaCl range (%, w/v)

11

2.5

18

Doubling timec (h)

nd, not determined; TP, trypticase peptones; YE, yeast extract; LC, liquid chromatography; BD, buoyant density method; Tm, melting point method a Environment from which the type strain was isolated b Substrates in parentheses are oxidized, but do not result in growth c Doubling time of the type strain under optimal growth conditions of temperature, pH, and NaCl d Cells grew in vitamin-free medium containing acetate

Type strain

Cell width (mm)

Reference Zellner et al. (1988)

29.4 (Tm) Koenig (1984)

54.5 (Tm) Kotelnikova et al. (1998)

34 (Tm)

GC content (mol%)

43

. Table 1 (Continued)

562 Methanobacteriales

RFM-2

RFM-1

RFM-3

HO

ZA-10

KM1H5– Ovine 1P rumen

ZR

curvatus

cuticularis

filiformis

gottschalkii

millerae

olleyae

oralis

0.7

0.23– 0.28

0.4

0.34

0.5

0.3– 0.5

Human subgingival plaque

0.4– 0.5

nd

Bovine rumen nd

Horse feces

Termite hindgut

Termite hindgut

Termite hindgut

Decaying cottonwood tissue

DH1

arboriphilus

Acidogenic digestor

ATM

Sourcea

acididurans

Species

Type strain

Cell width (mm) Methanogenesis substrates

H2 + CO2, formate

H2 + CO2, formate

H2 + CO2

H2 + CO2

H2 + CO2 (formate)d

H2 + CO2

0.7–1.2 H2 + CO2

nd

nd

0.9

4

1.2

1.6

1.2–1.4 H2 + CO2 (formate)c

0.3–0.5 H2 + CO2

Cell length (mm)

6.5–8.5 (7.7)

10- < 42 (37)

28–42 (36–40)

33–43 (36–42)

27–41 (37)

6.2–8.0 (6.9–7.4)

6.0–10.0 (7.5)

5.5–10.0 (7.0–8.0)

5.0–10.0 (7)

6.0–7.5 (7.0–7.2)

6.5–8.5 (7.1–7.2)

10- < 37 (30)

10–33.5 (30)

6.0–8.6 (7.5–8)

5.0–7.5 (6.0)

25–45 (30–37)

25–37 (35)

nd

nd

nd

37

35

40

13

16

0.06–0.6 15

up to 2.6

up to 2.6

nd

nd

nd

nd

0–0.6

nd

Temperature NaCl range pH range range Doubling (optimum) (optimum) (%, w/v) timeb (h) (oC)

Fecal extract 25–39 (35–38)

ac

ac, YE or TP

ac or YE or TP

YE

None

RF

B-vit

RF, ac, AAs

Required organic compounds

. Table 2 Descriptive characteristics of the species of the genus Methanobrevibacter

28 (Tm)

27–29 (Tm)

31–32 (Tm)

29 (Tm)

nd

nd

nd

25.5– 31.6 (Bd or Tm)

nd

GC content (mol%)

Ferrari et al. (1994)

Rea et al. (2007)

Rea et al. (2007)

Miller and Lin (2002)

Leadbetter et al. (1998)

Leadbetter and Breznak (1996)

Leadbetter and Breznak (1996)

Zeikus and Henning (1975)

Savant et al. (2002)

Reference

Methanobacteriales

43 563

CW

GS

SH

thaueri

woesei

wolinii

0.6

0.6

0.5

Methanogenesis substrates

H2 + CO2 (formate)d

H2 + CO2 (formate)d

1.0–1.4 H2 + CO2

1

0.6–1.2 H2 + CO2

1

0.8–1.7 H2 + CO2 (formate)d

Cell length (mm)

ac or YE or TP

ac or YE or TP

ac or YE or TP

ac, B-vit

nd (37)

nd (37)

nd (37)

26–46 (34–46)

nd (7)

nd (7)

nd (7)

5.0–8.5 (5.5–7.0)

5.5–7.7 (6–7)

nd

nd

nd

nd

nd

nd

nd

nd

nd

nd

Temperature NaCl range pH range range Doubling (optimum) (optimum) (%, w/v) timeb (h) (oC)

ac, B-vit, 33–42 CoM, 2-MBA, (37–39) AAs

Required organic compounds

Reference

33 (Tm)

31 (Tm)

38 (Tm)

30–31 (Tm or Bd)

Miller and Lin (2002)

Miller and Lin (2002)

Miller and Lin (2002)

Balch et al. (1979)

30.6 (Bd) Smith and Hungate (1958)

GC content (mol%)

nd, not determined; RF, rumen fluid; ac, acetate; AAs, amino acids; B-vit, B-vitamins; TP, trypticase peptones; YE, yeast extract; CoM, 2-mercaptoethanesulfonic acid (coenzyme M); 2-MBA, 2-methylbutyric acid; BD, buoyant density method; Tm, melting point method a Environment from which the type strain was isolated b Doubling time of the type strain under optimal growth conditions of temperature, pH, and NaCl c Formate is used by some, but not all strains d Growth on formate is poor

Sheep feces

Goose feces

Cow feces

Sewage sludge

PS

smithii

0.6– 0.7

Bovine rumen 0.7

Sourcea

ruminantium M1

Species

Cell width (mm)

43

Type strain

. Table 2 (Continued)

564 Methanobacteriales

1R7

Species

cuniculi

1

0.6–1.2

Cell width (mm)

1

0.6–1.2

Cell length (mm)

H2 + methanol

H2 + methanol

Methanogenesis substrates

Thiamine, ac, Ile, Leu

ac

Required organic compounds

nd, not determined; ac, acetate; Ile, isoleucine; Leu, leucine; Tm, melting point method a Environment from which the type strain was isolated

stadtmanae MCB3 Human feces

Rabbit rectum

Type strain Sourcea

. Table 3 Descriptive characteristics of the species of the genus Methanosphaera

30–40 (36–40)

>25– < 45 (35– 40)

Temperature range (optimum) (oC)

nd (6.5– 6.9)

nd (6.8)

pH range (optimum)

nd

nd

NaCl range (%, w/v)

25.8 (Tm)

23 (Tm)

GC content (mol%)

Miller and Wolin (1985)

Biavati et al. (1988)

Reference

Methanobacteriales

43 565

DSM2970 Sewage sludge and river sediment

wolfeii

H2 + CO2, formate

H2 + CO2 (formate)c

H2 + CO2

H2 + CO2, formate

2.5–6

H2 + CO2, formate

1.4–6.5 H2 + CO2

7–20

3–7

3–6

3–6

None

CoM

CoM

None

None

CoM

nd, not determined; CoM, 2-mercaptoethanesulfonic acid (coenzyme M); Tm, melting point method a Environment from which the type strain was isolated b Doubling time of the type strain under optimal growth conditions of temperature, pH, and NaCl c Formate is used by some, but not all strains

0.4– 0.6

0.36

M

thermophilus

Sludge of methane tank

Anaerobic 0.4 digestor

IDZ

0.35– 0.6

0.4– 0.6

thermoplexus

Sewage sludge

Sewage sludge

Marburg

marburgenesis

Anaerobic 0.4 digestor

Sourcea

thermoautotrophicus DH

ADZ

defluvii

Species

Type strain

Required Methanogenesis organic substrates compounds

37–74 (55–65)

47–75 (57)

45–70 (55)

40–75 (65–70)

45–70 (65)

45–65 (60)

6.0–8.2 (7.0–7.5)

6.5–8.5 (7.5)

7.5–8.5 (7.9–8.2)

6.0–8.8 (7.2–7.6)

5.0–8.0 (6.8–7.4)

6.0–7.5 (7.0)

2–3

3.5

3

1.6–2.5

nd (up 3.5–4 to 1)

0–0.6

0.1–3

0.01– 3.5

0.01– 3.5

Reference

Kotelnikova et al. (1993)

Scho¨nheit et al. (1980); Zeikus and Wolee (1972)

61 (Tm)

Winter et al. (1984)

44.7 (Tm) Laurinavichyus et al. (1988)

55 (Tm)

49 (Tm)

47.6 (Tm) Wasserfallen et al. (2000)

62.2 (Tm) Kotelnikova et al. (1993)

GC Doubling content timeb (h) (mol%)

0.08–2 1.5

NaCl Temperature range range pH range (%, (optimum) (optimum) w/v) (oC)

43

Cell Cell width length (mm) (mm)

. Table 4 Descriptive characteristics of the species of the genus Methanothermobacter

566 Methanobacteriales

Icelandic hot spring

fervidus

sociabilis Kf1F1

0.3– 0.4

0.3– 0.4

Cell width (mm)

1–3

1–3

Cell length (mm)

H2 + CO2

H2 + CO2

Methanogenesis substrates

None

None

Required organic compounds

55–97 (88)

67–97 (80–85)

5.5–7.5 (6.5)

nd (6.5)

nd

nd

3

3

NaCl Temperature pH range range Doubling range (optimum) (oC) (optimum) (%, w/v) timeb (h)

nd, not determined; Tm, melting point method a Environment from which the type strain was isolated b Doubling time of the type strain under optimal growth conditions of temperature, pH, and NaCl

Icelandic hot spring

V24S

Species

Sourcea

Type strain

. Table 5 Descriptive characteristics of the species of the genus Methanothermus

33 (Tm)

33 (Tm)

Lauerer et al. (1986)

Stetter et al. (1981)

GC content (mol%) Reference

Methanobacteriales

43 567

568

43

Methanobacteriales

The placement of the hyperthermophilic Methanothermus into a separate family from other Methanobacteriales genera is justified by the deep branching of the phylogeny of its 16S rRNA gene (> Fig. 1). The 16S rRNA gene sequence similarities within Methanothermus species are much higher (98%) than similarities between Methanothermus

. Figure 1 Phylogenetic tree for the Methanobacteriales based upon 16S rRNA sequences. The alignment was manually edited to include 1,308 positions. The 16S rRNA sequences of Methanocaldococcus jannaschii (M59126) and Methanococcus vannielii (AY196675) were used as outgroups to generate the root. The tree was constructed with the neighbor-joining algorithm in MEGA4. Bootstrap analysis was performed with 1,000 replicates, and values greater than 50% are labeled on the nodes. The scale bar is 0.02 expected nucleotide substitutions per site. The GenBank accession numbers for the sequences is given in parentheses following their names.

Methanobacteriales

43

and other members of the Methanobacteriales (83–89%). This classification is further confirmed by DNA reassociation. For instance, the DNA relatedness between Methanothermus isolates and Methanothermobacter thermoautotrophicus strain IM is 2–8% (Lauerer et al., 1986). Phenotypically, the genus Methanothermus is distinguished from other Methanobacteriales by their high temperature optima (80–88 C), double-layered cell wall, and motility by bipolar polytrichous flagellation. Methanobacteriaceae is a diverse family, including mesophilic and thermophilic species. The phylogeny of the 16S rRNA gene indicates that the thermophilic species are divergent from mesophilic members at the genus level. The 16S rRNA sequence similarities within the thermophilic genus Methanothermobacter are above 98%, while the similarities between thermophilic and mesophilic members of Methanobacteriaceae are generally below 93% (Wasserfallen et al., 2000). The DNA relatedness between Methanothermobacter species are 22–47%, confirming that they are genetically distant and should be assigned to separate species (Boone et al., 2001). The separation of mesophilic members of Methanobacteriales into three genera is supported by both genetic and phenotypic analyses. Species of Methanobacterium are usually autotrophs, while species of Methanobrevibacter and Methanosphaera are commonly mixotrophs or heterotrophs. Species of Methanosphaera use only H2 and methanol as substrates for methanogenesis, while all species of Methanobrevibacter and Methanobacterium can use H2 and CO2.

4

Ecology

Members of the Methanobacteriales are widely distributed in anaerobic habitats such as marine and freshwater sediments, soils, animal gastrointestinal tracts, anaerobic sewage digestors, and geothermal habitats. Methanobacterium has been cultivated from marine and freshwater sediments, groundwaters, soils, anaerobic digestors, and animal gastrointestinal tracts and has also been detected as endosymbionts in anaerobic ciliates (Embley et al., 1992). Methanobrevibacter has been isolated from rumens, feces, termite hindguts, human subgingival plaque, anaerobic digestors, and decaying wood tissues. Methanosphaera has only been isolated from animal gastrointestinal tracts but has been detected in anaerobic digestors (Weiss et al., 2008). Methanothermobacter has been cultivated from thermophilic anaerobic digestors and natural gas and oil fields (Nazina et al., 2006; Mochimaru et al., 2007). Methanothermus has only been isolated from solfarata hot springs.

5

Research Needs

Our current knowledge on the diversity of the Methanobacteriales is largely incomplete. As an example, investigations of 16S rRNA gene from clone libraries recognized a large number of uncultured Methanobrevibacter, especially from the rumen and termite gut (Dighe et al., 2004; Wright et al., 2004). Moreover, the cloned sequences from termite gut formed separate lineages from cultured Methanobrevibacter (Dighe et al., 2004). Any correlations between habitat and 16S rRNA based phylogeny need more ecological surveys to unravel.

569

570

43

Methanobacteriales

References Balch WE, Fox GE, Magrum LJ, Woese CR, Wolfe RS (1979) Methanogens: reevaluation of a unique biological group. Microbiol Mol Biol Rev 43: 260–296. Belyaev SS, Wolkin R, Kenealy WR, Deniro MJ, Epstein S, Zeikus JG (1983) Methanogenic bacteria from the Bondyuzhskoe oil field: general characterization and analysis of stable-carbon isotopic fractionation. Appl Environ Microbiol 45: 691–697. Biavati B, Vasta M, Ferry JG (1988) Isolation and characterization of ‘‘Methanosphaera cuniculi’’ sp. nov. Appl Environ Microbiol 54: 768–771. Bonin A, Boone D (2006) The order Methanobacteriales. In The Prokaryotes, 3rd edn. M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.). New York: Springer-Verlag, pp. 231–243. Boone DR (1987) Request for an opinion: replacement of the type strain of Methanobacterium formicicum and reinstatement of Methanobacterium bryantii sp. nov. nom. rev. (ex Balch and Wolfe, 1981) with M.o.H. (DSM 863) as the type strain. Int J Syst Bacteriol 37: 172–173. Boone DR, Whitman WB, Koga Y (2001) Methanobacteriales. In Bergy’s Manual of Systematic Bacteriology, 2nd edn. DR Boone, RW Castenholtz, GM Garrity (eds.). New York: Springer-Verlag, pp. 213–235. Cuzin N, Ouattara AS, Labat M, Garcia JL (2001) Methanobacterium congolense sp. nov., from a methanogenic fermentation of cassava peel. Int J Syst Evol Microbiol 51: 489–493. Dighe A, Jangid K, Gonzalez J, Pidiyar V, Patole M, Ranade D, Shouche Y (2004) Comparison of 16S rRNA gene sequences of genus Methanobrevibacter. BMC Microbiol 4: 20. Embley TM, Finlay BJ, Thomas RH, Dyal PL (1992) The use of rRNA sequences and fluorescent probes to investigate the phylogenetic positions of the anaerobic ciliate Metopus palaeformis and its archaeobacterial endosymbiont. J Gen Microbiol 138: 1479–1487. Ferrari A, Brusa T, Rutili A, Canzi E, Biavati B (1994) Isolation and characterization of Methanobrevibacter oralis sp. nov. Curr Microbiol 29: 7–12. Joulian C, Patel BKC, Ollivier B, Garcia JL, Roger PA (2000) Methanobacterium oryzae sp. nov., a novel methanogenic rod isolated from a Philippines ricefield. Int J Syst Evol Microbiol 50: 525–528. Koenig H (1984) Isolation and characterization of Methanobacterium uliginosum sp.nov. from a marshy soil. Can J Microbiol 30: 1477–1481. Kotelnikova S, Macario AJL, Pedersen K (1998) Methanobacterium subterraneum sp. nov., a new

alkaliphilic, eurythermic and halotolerant methanogen isolated from deep granitic groundwater. Int J Syst Bacteriol 48: 357–367. Kotelnikova SV, Obraztsova AY, Gongadze GM, Laurinavichius KS (1993) Methanobacterium thermoflexum sp. nov. and Methanobacterium defluvii sp. nov.: Thermophilic rod-shaped methanogens isolated from anaerobic digestor sludge. Syst Appl Microbiol 16: 427–435. Lauerer G, Kristjansson JK, Langworthy TA, Koenig H, Stetter KO (1986) Methanothermus sociabilis sp. nov., a second species within the Methanothermaceae growing at 97oC. Syst Appl Microbiol 8: 100–105. Laurinavichyus KS, Kotel’nikova SV, Obraztsova AY (1988) New species of thermophilic methaneproducing bacteria Methanobacterium thermophilum. Mikrobiologiya 57: 1035–1041. Leadbetter JR, Breznak JA (1996) Physiological ecology of Methanobrevibacter cuticularis sp. nov. and Methanobrevibacter curvatus sp. nov., isolated from the hindgut of the termite Reticulitermes flavipes. Appl Environ Microbiol 62: 3620–3631. Leadbetter JR, Crosby LD, Breznak JA (1998) Methanobrevibacter filiformis sp. nov., a filamentous methanogen from termite hindguts. Arch Microbiol 169: 287–292. Ma K, Liu X, Dong X (2005) Methanobacterium beijingense sp. nov., a novel methanogen isolated from anaerobic digesters. Int J Syst Evol Microbiol 55: 325–329. Miller TL, Lin C (2002) Description of Methanobrevibacter gottschalkii sp. nov., Methanobrevibacter thaueri sp. nov., Methanobrevibacter woesei sp. nov. and Methanobrevibacter wolinii sp. nov. Int J Syst Evol Microbiol 52: 819–822. Miller TL, Wolin MJ (1985) Methanosphaera stadtmaniae gen. nov., sp. nov.: a species that forms methane by reducing methanol with hydrogen. Arch Microbiol 141: 116–122. Mochimaru H, Yoshioka H, Tamaki H, Nakamura K, Kaneko N, Sakata S, Imachi H, Sekiguchi Y, Uchiyama H, Kamagata Y (2007) Microbial diversity and methanogenic potential in a high temperature natural gas field in Japan. Extremophiles 11: 453–461. Nazina T, Shestakova N, Grigor’yan A, Mikhailova E, Tourova T, Poltaraus A, Feng C, Ni F, Belyaev S (2006) Phylogenetic diversity and activity of anaerobic microorganisms of high-temperature horizons of the Dagang oil field (P. R. China). Microbiology 75: 55–65. Patel GB, Sprott GD, Fein JE (1990) Isolation and characterization of Methanobacterium espanolae sp. nov.,

Methanobacteriales a mesophilic, moderately acidiphilic methanogen. Int J Syst Bacteriol 40: 12–18. Rea S, Bowman JP, Popovski S, Pimm C, Wright A-DG (2007) Methanobrevibacter millerae sp. nov. and Methanobrevibacter olleyae sp. nov., methanogens from the ovine and bovine rumen that can utilize formate for growth. Int J Syst Evol Microbiol 57: 450–456. Savant DV, Shouche YS, Prakash S, Ranade DR (2002) Methanobrevibacter acididurans sp. nov., a novel methanogen from a sour anaerobic digester. Int J Syst Evol Microbiol 52: 1081–1087. Scho¨nheit P, Moll J, Thauer RK (1980) Growth parameters (Ks, mmax, Ys) of Methanobacterium thermoautotrophicum. Arch Microbiol 127: 59–65. Shlimon AG, Friedrich MW, Niemann H, Ramsing NB, Finster K (2004) Methanobacterium aarhusense sp. nov., a novel methanogen isolated from a marine sediment (Aarhus Bay, Denmark). Int J Syst Evol Microbiol 54: 759–763. Smith PH, Hungate RE (1958) Isolation and characterization of Methanobacterium ruminantium N. SP. J Bacteriol 75: 713–718. Stetter KO, Thomm M, Winter J, Wildgruber G, Huber H, Zillig W, Janecovic D, Koenig H, Palm P, Wunderl S (1981) Methanothermus fervidus, sp. nov., a novel extremely thermophilic methanogen isolated from an Icelandic hot spring. Mikrobiol Hyg, I Abt C 2: 166–178. Wasserfallen A, Nolling J, Pfister P, Reeve J, Conway de Macario E (2000) Phylogenetic analysis of 18 thermophilic Methanobacterium isolates supports the proposals to create a new genus, Methanothermobacter gen. nov., and to reclassify several isolates in

43

three species, Methanothermobacter thermautotrophicus comb. nov., Methanothermobacter wolfeii comb. nov., and Methanothermobacter marburgensis sp. nov. Int J Syst Evol Microbiol 50: 43–53. Weiss A, Je´roˆme V, Freitag R, Mayer H (2008) Diversity of the resident microbiota in a thermophilic municipal biogas plant. Appl Microbiol Biotechnol 81: 163–173. Winter J, Lerp C, Zabel H-P, Wildenauer FX, Koenig H, Schindler F (1984) Methanobacterium wolfei, sp. nov., a new tungsten-requiring, thermophilic, autotrophic methanogen. Syst Appl Microbiol 5: 457–466. Worakit S, Boone DR, Mah RA, Abdel-Samie M-E, El-Halwagi MM (1986) Methanobacterium alcaliphilum sp. nov., an H2-utilizing methanogen that grows at high pH values. Int J Syst Bacteriol 36: 380–382. Wright A-DG, Williams AJ, Winder B, Christophersen CT, Rodgers SL, Smith KD (2004) Molecular diversity of rumen methanogens from sheep in western Australia. Appl Environ Microbiol 70: 1263–1270. Zeikus JG, Henning DL (1975) Methanobacterium arbophilicum sp.nov. An obligate anaerobe isolated from wetwood of living trees. Antonie Van Leeuwenhoek 41: 543–552. Zeikus JG, Wolee RS (1972) Methanobacterium thermoautotrophicus sp. n., an anaerobic, autotrophic, extreme thermophile. J Bacteriol 109: 707–713. Zellner G, Bleicher K, Braun E, Kneifel H, Tindall BJ, Macario EC, Winter J (1988) Characterization of a new mesophilic, secondary alcohol-utilizing methanogen, Methanobacterium palustre spec. nov. from a peat bog. Arch Microbiol 151: 1–9.

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44 Methanococcales Y. Liu Department of Microbiology, University of Georgia, Athens, GA, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 574 2 Descriptive Features . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 574 3 Phylogeny and Taxonomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 574 4 Ecology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 580 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 580

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_44, # Springer-Verlag Berlin Heidelberg, 2010

574

44

Methanococcales

Abstract: The order Methanococcales is composed of two families, Methanocaldococcaceae and Methanococcaceae, which are distinguished by 16S rRNA sequence similarities below 93% and differences in growth temperatures. The Methanocaldococcaceae are all hyperthermophilic, while the Methanococcaceae are extremely thermophilic or mesophilic. Members of this order are all capable of forming methane by CO2 reduction with H2. Many species can use formate as an alternative electron donor. Most species can grow autotrophically.

1

Introduction

Members of the Methanococcales (the methanococci) are coccoid methanogens isolated from marine environments. They share a set of phenotypic characteristics. They all use H2 or formate to reduce CO2 for methanogenesis. Acetate, methyl-containing compounds, and alcohols are not used as substrates for methanogenesis. Most of them can grow autotrophically with CO2 as the sole carbon source. Sulfide is a sufficient sulfur source for all methanococci, and elemental sulfur is reduced to sulfide with slight inhibition of growth in most strains. Ammonium is a suitable nitrogen source for all methanococci, and nitrogen gas, nitrate, and alanine are used as a nitrogen source by some species. They all require sea salts for optimal growth. Cells are irregular cocci, 1–3 mm in diameter during balanced growth. Most of them are motile by means of polar tuft(s) of flagella. Cells strain Gram-negative. They are susceptible to lysis by 0.01% (w/v) SDS and hypotonic solutions. Cell envelopes are composed of a protein cell wall or S-layer. Glycoproteins and cell wall carbohydrates are not abundant. The cellular lipids contain archaeol, caldarchaeol, hydroxyarchaeol, and macrocyclic archaeol, depending upon the species. The polar lipids can contain glucose, N-acetylglucosamine, serine, and ethanolamine. The optimal growth temperatures of methanococci are diverse, ranging from 35 to 88 C. They are among the fastest growing methanogens at either moderate or high temperatures, with generation times of about 2 h at 37 C and less than 30 min at 85 C.

2

Descriptive Features

Descriptive properties of the methanococci are summarized in > Tables 1 and 2. Further information can be found in (Whitman et al., 2001; Whitman and Jeanthon, 2006).

3

Phylogeny and Taxonomy

Phylogenetic analyses with DNA sequences reveal a high diversity of the Methanococcales. The sequence similarities of the 16S rRNA genes between hyperthermophilic and mesophilic methanococci are generally below 90%. For instance, the 16S rRNA gene sequence similarity between the mesophile Methanococcus voltae and hyperthermophile Methanocaldococcus infernus is about 85%, which is comparable to the similarity between Escherichia and Pseudomonas. In addition, the mesophilic methanococci possess 91–96% (average 94%) 16S rRNA gene sequence similarities and 5–30% DNA reassociation values, suggesting that they are related only at the genus level (Keswani et al., 1996). The Methanococcales are currently divided into two families and four genera, according to their growth temperatures. The family Methanocaldococcaceae includes two hyperthermophilic

H2 + CO2

H2 + CO2

+

Substrates for methanogenesis

Autotrophy

5.2–7.0

6.0

pH range

pH optimum

NaCl range (%, w/v) 1.0–5.0

85

85

Temperature optimum ( C)

0.8–3.5

6.5

5.25–7.0

55–91

50–91

Temperature range ( C)

S ,S

S ,S

Sulfur source

0

0.5–5.0

6.5

5.5–7.6

85

48–92

0

S ,S

2

NH3, NO3

2

2

NH3, NO3

Nitrogen source

NH3

+

+

Selenium simulates + growth

+

+

+

+

H2 + CO2

nd

1–2

AG86

fervens

indicus

1.5–5.0

6.5

5.5–6.7

85

50–86

0

S ,S

2

NH3, NO3

+

+

+

H2 + CO2

1 tuft

1–3

SL 43

Methanocaldococcus

Yeast extract stimulates growth

0

3 tufts

2 tufts

Flagellaa

1–3

1.5

Cell diameter (mm)

ME

infernus

JAL-1

jannaschii

Type strain

Character

0.6–5.6

6.5

5.2–7.0

80

49–89

0

S ,S

2

NH3, NO3

+

+

+

H2 + CO2

3 tufts

1–3

M7

vulcanius

. Table 1 Descriptive characteristics of the species of the genera Methanocaldococcus and Methanotorris

formicicus

0

0.9–5.4

5.7

5.0–7.5

88

45–91

S ,S

2

NH3

+

0.4–6.0

6.7

6.0–8.5

75

55–83

S2

NH3, N2, NO3

+

H2 + CO2, formate



H2 + CO2

0.8–1.5



Mc-S-70

Methanotorris

1–2

Kol 5

igneus

Methanococcales

44 575

Jones et al. (1983a)

Reference

Jeanthon et al. (1999) Zhao et al. (1988)

L’Haridon et al. (2003)

25–30

30

31 (Tm)

1.8

igneus

Jeanthon et al. (1999)

Burggraf et al. (1990)

formicicus

Takai et al. (2004)

Deep-sea black smoker chimney

30

33 (LC)

2.4

Methanotorris

Deep-sea Shallow marine hydrothermal hydrothermal vent vent

45

31 (Tm)

2.5

vulcanius

nd, not determined; Bd, buoyant density method; Tm, melting point method; LC, liquid chromatography a Number of flagellar tufts. , non-motile, but flagella-like structures are observed by electron microscopy b Doubling time of the type strain under optimal growth conditions of temperature, pH, and NaCl c Environment from which the type strain was isolated

Jeanthon et al. (1998)

Deep-sea Deep-sea Deep-sea Deep-sea hydrothermal hydrothermal hydrothermal hydrothermal vent vent vent vent

Sourcec

20–30

35–40

26

Doubling time (min)b

31 (LC)

33 (Tm)

33 (Tm)

3.0

indicus

GC content (mol%) 31 (Bd)

fervens

Methanocaldococcus

3.0

3.0

infernus

2.0

NaCl optimum (%, w/v)

jannaschii

44

Character

. Table 1 (Continued)

576 Methanococcales

Multiple tufts

H2 + CO2, formate

2 tufts

H2 + CO2, formate

+

Flagellaa

Substrates for methanogenesis

Autotrophy

10– < 55 (45)

22–50 (40)

10–45 (20–25)

15–40 (25–30)

37–45 (35–40)

Temperature range (optimum) (oC)

6.2–7.8 (6.5–7.2)

5.0–8.7 (6.0–7.5)

6.5–8.0 (6.9–7.5)

5.8–7.6 (6.2–6.6)

6.7–8.0 (6.7–7.2)

5.5–8.0 (6.7)

pH range (optimum)

1.2

0.6–2.3

nd

0.6–1.1

0.6

0.2–1

NaCl optimum (%, w/v)

2.5

6.8

13.5

10

46

18

Doubling timec (h)

Ollivier et al. (1986)

Reference

Romesser et al. (1979)

55–60 (Tm)

nd

Rivard and Smith (1982)

Mikucki et al. (2003)

59.5 (LC) Zellner et al. (1998)

61 (Bd)

62.2 (LC) Dianou et al. (2001)

59 (Bd)

GC content (mol %)

Methanomicrobiales

45 585

ML15

GKZPZ

Chile 9

formosanus

liminatans

tationis

JR1

Ace-2

cariaci

frigidum

Methanogenium

N2F9704

aquaemaris

Methanofollis

Organism

Ø 1.5–2.0

Ø 1.2–2.0

Anoxic Ace Lake water

Marine sediments

Solfataric pool mud

Ø 1.5–2.5

Ø < 2.6

Ø 1.5–3

Wastewater Ø 1.25–2.0 reactor

Marine– water fish pond

Marine– water fish pond

Sourcea

Dimensions (mm)

None

Pertrichous

Peritrichousf

Flagellatedf

None

None

Flagella

H2 + CO2, formate

H2 + CO2, formate

H2 + CO2, formate

H2 + CO2, formate, 2-propanol, 2-butanol, cyclopentanol

H2 + CO2, formate

H2 + CO2, formate

Methanogenesis substratesb

ac

ac, YE

ac, YE, TP, tung

ac

YE, TP

None

Required organic compounds

12g-18 (15)

10–32 (20–25)

6.5–7.9 (7.5–7.9)

nd (6.8– 7.3)

6.3–8.8 (7)

nd (7)

15–44 (40)

15–44 (40–44)

5.6–7.3 (6.6–7.0)

6.3–8.0 (6.5)

pH range (optimum)

20–42 (40)

20–43 (37)

Temperature range (optimum) (oC)

2–3.5

2.7

0.8–1.2

0–3.5

3

0.5

NaCl optimum (%, w/v)

69.6

11

12

7.5

36

13

Doubling timec (h)

Reference

nd

52 (Bd)

54 (Tm)

60 (Tm)

Franzmann et al. (1997)

Romesser et al. (1979)

Zabel et al. (1984)

Zellner et al. (1990)

58.4 (Tm) Wu et al. (2005)

59.1 (Tm) Lai and Chen (2001)

GC content (mol %)

45

Type strain

. Table 1 (Continued)

586 Methanomicrobiales

SEBR4847 Offshore oil Ø 1–3 field

0.1– 0.31.5–2.8

petrolearius

0.5–11.6– 3.4

M3

Swamp

Flagellatedd

None

Flagellatedd

None

Polar tuft

peritrichous

0.7 1.5–2.0 Single

0.61.5–2.5

limicola

Marine ciliate

Bovine rumen

Marine sediment

MC1

BP

Ø 1–1.2

Marine mud Ø 0.5–1.5

Marine sediments

endosymbiosus

Methanoplanus

mobile

Methanomicrobium

paynteri

G2000

CV

organophilum

Methanolacinia

AK-1

marinum

H2 + CO2, formate, 2-propanol

H2 + CO2, formate

H2 + CO2, formate

H2 + CO2, formate

H2 + CO2, 2-propanol, 2-butanol

H2 + CO2, formate, 2-propanol, 2-butanol, ethanol

H2 + CO2, formate

5–25 (25)

ac

ac

p-Cresol, tung

Complex

ac

28–43 (35–40)

17–41 (40)

16–36 (32)

35–45 (40)

20–45 (40)

ac, PABA, nd-39 (30–35) biotin, tung, vit-B

ac

5.3–8.2 (7.0)

nd (6.5– 7.5)

6.1–8.0 (6.8–7.3)

5.9–7.7 (6.1–6.9)

6.6–7.3 (7.0)

nd (6.4– 7.3)

5.5–7.5 (6.0)

1–3

1

1.5

nd

0.88

2.0

1.5–7.3

10

7

7

nd

4.8

6

42

Chong et al. (2002)

50 (LC)

Ollivier et al. (1997)

47.5 (Tm) Wildgruber et al. (1982)

38.7 (Tm) Bruggen et al. (1986)

48.8 (Bd) Paynter and Hungate (1968)

44.9 (Bd) Rivard et al. (1983)

46.7 (Tm) Widdel et al. (1988)

nd

Methanomicrobiales

45 587

JF-1

Z

XII

China Z

labreanum

parvum

sinense

chunghsingensis

Pilot plant for wastewater treatment

Anaerobic digestor

Lake sediments

Sediment of wastewater treatment pond

Sewage sludge

Sourcea

K1F9705b Marine water fishpond

Methanocalculus

SZSXXZ

bavaricum

Methanocorpusculum

hungatei

Methanospirillum

Organism

Flagellatede

Flagellated

Ø FeOOH > SO42
> CO2, and in general the most favored electron acceptor will be used preferentially. Methanogenesis from the reduction of CO2 with hydrogen is the least energetically favorable mode of anaerobic respiration among all mentioned above. In marine

Introduction

49

environments, sulfate-reducing bacteria consume the majority of available H2 and acetate (Capone and Kiene, 1988). Because of the competitive relationships between different microorganisms for common substrates, significant methanogenesis occurs only when more favorable electron acceptors are absent or exhausted. Besides methanogens, homoacetogens are another group of anaerobes that can reduce CO2 for energy production (Drake et al., 2008). Acetogenesis with H2 and CO2 is thermodynamically less favorable than methanogenesis. Therefore, homoacetogens do not compete well with methanogens in many habitats. However, homoacetogens outcompete methanogens in some environments, such as the hindgut of certain termites and cockroaches. Unlike methanogens they can consume some sugars and be involved in the demethylation of some low molecularweight compounds. Low temperature can also provide a selective advantage to homoacetogens, which reduce CO2 to acetate instead of methane (Kotsyurbenko, 2005). This limits H2-dependent methanogenesis and favors to acetoclastic methanogenic pathway. A similar effect was also observed in slightly acidic lake sediments (Casper et al., 2003). Moreover, H2 microbial production becomes less favorable and decreases at low temperature. Therefore, H2-dependent methanogenesis in cold environments can be inhibited due to an insufficient supply of substrates. H2-consuming methanogens can be also inhibited by decreasing in pH or in the presence of toxic compounds. It results in the increase of the pools of fatty acids that acidifies the whole system and stops methanogenesis (Hori et al., 2006). In opposite, hydrogen-dependent methanogens predominate in ruminants where fatty acids produced in the rumen during cellulose degradation are then rapidly absorbed by the gut as an energy source whereas H2 and CO2 are consumed by methanogens. Thus, changes in environmental conditions can cause different responses of constituent microorganisms, rearrange their trophic relations and affect the functioning of the entire community. It in turn can result then in redistribution of main flows of organic matter in the community and affect the methanogenic pathways (Conrad, 2002; Kotsyurbenko, 2005; Kotsyurbenko et al., 2007). In sum, the anaerobic community represents a biological system that is balanced by the coordinated interactions of the constituent microbial groups. The methane biocenosis is able to change its fermentative pathways and to function as a self-regulatory system. Methanogens act as the primary regulators in the trophic microbial interactions at the anaerobic degradation processes.

3

Methods to Study Methanogenic Communities

The investigation of microbial communities active in different ecosystems basically includes two main approaches. The study of the community as a whole operating under as close to the in situ conditions as possible and assess their contribution to natural processes, minimally invasive methods are required in order to identify their spatial distributions and relate these to activity. Alternatively, the study of the trophic structure and constituent functional groups of microorganisms as interacting components requires a deliberate imbalance of the community. The former is mainly based on biogeochemical investigations involving measuring actual methane production in situ and in laboratory experiments as well as using different radioactive tracers as the immediate methanogenic precursors that can be incorporated into CH4 (Chidthaisong et al., 1999; Schulz et al., 1997) and isotopic fractionation during

621

622

49

Introduction

methanogenesis (Fey et al., 2004) under steady state conditions. Molecular biological and biochemical methods can be applied to study the qualitative and quantitative composition of different microbial groups in the community (Friedrich, 2006; Kirka et al., 2004; Lueders et al., 2004; Spiegelman et al., 2005). The latter can be achieved by diluting the system and/or introducing the specific substrates of key microorganisms and metabolic inhibitors into the community (Conrad and Klose, 2000; Kotsyurbenko et al., 1993). The combined use of various molecular techniques, for example T-RFLP, DGGE, 16S rRNA gene cloning, real-time PCR and microscopic evidence (FISH), provides valuable tools to get insight into ecophysiology of the complex methanogenic communities (Amann et al., 1995; Collins et al., 2006; Yu et al., 2006). Methanogens, the key microbial group of the methanogenic community belong to the kingdom of Euryarchaeota in the domain of Archaea (Liu and Whitman, 2008). They can be separated into three main nutritional categories: hydrogenotrophs (oxidizing H2 and CO2), methylotrophs (utilizing methyl compounds as methanol, methylamines or dimethylsulfides) and acetoclastic methanogens (utilizing the methyl group of acetate). Most methanogens are hydrogenotrophs and belong to the orders Methanobacteriales, Methanococcales, Methanomicrobiales and Methanopyrales. Many of them can also use formate as the major electron donor. Members of the order Methanosarcinales have the widest substrate range among methanogens including all three above mentioned substrate groups. Recent culture-independent studies have revealed the presence of novel phylogenetic groups of methanogens which are probably represent new orders (Erkel et al., 2006). The ability of methanogens to produce methane under anaerobic conditions has provided them with a unique set of enzymatic pathways, as well as certain coenzymes and cofactors that are not found in other organisms. Despite the isolation of the key methanogens operating in the ecosystem is important, traditional methods in microbiology, based on cultivation, can be hardly applied to make a comprehensive study of the diversity of methanogens in ecosystems. These archaea belong to organisms known to be difficult to cultivate. They are obligate anaerobes, some require long incubation periods, they are difficult to separate from their syntrophic partners, and require low reducing potential. Molecular biological methods combined with microscopical approaches including coenzyme F420 autofluorescence microscopy (Doddema and Vogels, 1978) are well suited for the study ecology of methanogenic archaea. The 16S-rRNA molecule is an excellent marker to infer phylogeny because it is found in all cellular life form. Primers for the study of methanogenic populations have been designed to amplify various regions of the 16S-rRNA. Some primers target the domain Archaea generally, other are specific for methanogenic 16S-rRNA (Hales et al., 1996; Upton et al., 2000). Another molecular marker widely used for studies of methanogenic diversity is the gene coding for methyl-coenzyme M reductase (MCR), a key enzyme for methanogenesis (Friedrich, 2005). The MCR catalyses the reduction of methyl-coenzyme M accompanied by release of methane. Unlike other enzymes in the methanogenic metabolism, the MCR appears to be unique to the methanogens (Thauer, 1998). Parts of the MCR operon are also highly conserved and all MCR operons appear to have evolved from a common source. This functional gene is therefore a helpful tool for an ecological and phylogenetic survey of a microbial population with known activity. Essential progress has been achieved in getting insights into the physiology of methanogens by genome sequencing. The completed methanogen genomes include by now 18 cultured strains (Liu and Whitman, 2008). Bionformatic, proteomic, microarray, and biochemical

Introduction

49

approaches investigating methanogens from different ecosystems are also useful to get insights into differences in their metabolism and adaptation capacities. Genomewide proteomic approach revealed novel enzymes and a mechanism for energy-conservation for Methanosarcina acetivorans, a methanogen isolated from marine sediments (Ferry and Lessner, 2008).

4

Research Needs

The anaerobic community represents a biological system that is balanced by the coordinated interactions of the constituent microbial groups. Such microbial interactions in various environments are complex. So, the study of biological production of methane should involve not only field measurements and isolation of the key microorganisms, but also microbial interactions at the community level. Although a link between methane production and different environmental parameters have been established in previous studies, there is a lack of information on how methanogenic decomposition pathways are related to microbial community structure. The important point is also changes in the trophic structure of the methanogenic community depending on different effects of environment and resulting in changes of the final products of its activity. To understand such changes and to resolve the linkage between structure and function in methanogenic communities, it is necessary to know which parts of its tropic chain is altered, what groups of microorganisms are especially influenced by a given factor, and what groups of microorganisms become determining in the community. Understanding the functionality of microbial communities in natural and engineered ecosystems is still a major objective for microbial ecologists.

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by anaerobic bacteria at low temperature. Chemosphere 27: 1745–1761. Lettinga G (1995) Anaerobic digestion and wastewater treatment systems. Antonie Van Leeuwenhoek 67: 3–28. Liu Y, Whitman WB (2008) Methabolic, phylogenetic, and ecological diversity of the metanogenic archaea. Ann N Y Acad Sci 1125: 171–189. Lueders T, Pommerenke B, Friedrich MW (2004) Stableisotope probing of microorganisms thriving at thermodynamic limits: syntrophic propionate oxidation in flooded soil. Appl Environ Microbiol 70: 5778–5786. Oremland RS, King GM (1989) Methanogenesis in hypersaline environments. In Microbial Mats: Physiological Ecology of Benthic Microbial Communities. Y Cohen and E Rosenberg (eds.). Washington DC: American Society for Microbiology, pp. 180–190. Pimenov NV, Ivanova AE (2005) Anaerobic methane oxidation and sulfate reduction in bacterial mats on coral-like carbonate structures in the Black Sea. Microbiology 74: 362–370. Reeburgh WS (2007) Oceanic methane biogeochemistry. Chem Rev 107: 486–513. Schink B, Stams AJM (2006) Syntrophism among Prokaryotes. In Prokaryotes vol. 2. M Dworkin, S Falkow, E Rosenberg, KH Schleifer, and E Stackebrandt (eds.). New York, Berlin: Springer Verlag, pp. 309–335. Schulz S, Matsuyama H, Conrad R (1997) Temperature dependence of methane production from different precursors in a profundal sediment (Lake Constance). FEMS Microbiol Ecol 16: 251–260. Spiegelman D, Whissell G, Greer CW (2005) A survey of the methods for the characterization of microbial consortia and communities. Can J Microbiol 51: 355–386 Thauer RK (1998) Biochemistry of methanogenesis: a tribute to Marjory Stephenson. Microbiology 144: 2377–2406. Upton M, Hill B, Edwards C, Saunders JR, Ritchie DA, Lloyd D (2000) Combined molecular ecological and confocal laser scanning microscopic analysis of peat bog methanogen populations. FEMS Microbiol Lett 193: 275–281. Whiticar MJ (1999) Carbon and hydrogen isotope systematics of bacterial formation and oxidation of methane. Chem Geol 161: 291–314. Yu Y, Kim J, Hwang S (2006) Use of real-time PCR for group-specific quantification of aceticlastic methanogens in anaerobic processes: population dynamics and community structures. Biotechnol Bioeng 93: 424–433. Zavarzin GA (1995) So¨hngen psychrophilic cycle. Ecol Chem 4: 3–12.

50 Soil, Wetlands, Peat O. R. Kotsyurbenko Technical University Braunschweig, Institute of Microbiology, Braunschweig, Germany [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 626 2 Anoxic Waterlogged Soils as a Habitat for the Methanogenic Community . . . . . . . . 627 3 Methanogenesis in Peatlands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 628 4 Methanogenic Diversity in Peatlands . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 630 5 Acidotolerant Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 631 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 631

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_50, # Springer-Verlag Berlin Heidelberg, 2010

626

50

Soil, Wetlands, Peat

Abstract: Soil is the naturally occurring rock particles and decaying organic matter (humus) on the surface of the earth, capable of supporting life. It has three components: solid, liquid, and gas. The solid phase is a mixture of mineral and organic matter. Wetlands are areas on which water covers the soil or where water is present either at or near the surface of that soil. Wetlands often host considerable biodiversity and endemism. Their hydrological conditions are characterized by an absence of free oxygen sometimes or always. It favors the development of anaerobic microbial community. In the absence of electron acceptors other than bicarbonate, methane is the end product of organic matter degradation in wetland ecosystems. It makes wetlands important sources of the greenhouse gas CH4 in the context of the problem of global climate changes. Peatlands are a type of wetlands and form when plant material is inhibited from decaying by acidic and anaerobic conditions. Methane production in peatlands tends to vary tremendously both spatially and temporally and depends on environmental factors such as temperature, pH, and water table, as well as plant cover. In anaerobic peat, acetate and CO2 are the most quantitatively important CH4 precursors. Most studies suggest that acetoclastic methanogenesis is an important pathway for CH4 formation in nutrient-rich fens covered with Carex sedges, whereas CO2 reduction is an important methanogenic pathway in Sphagnum-dominated bogs. Such bogs, the predominant peatlands, are typically acidic (pH < 5) with low concentrations of mineral nutrients. The Sphagnum bog microbes seem to have special metabolic mechanisms to cope with low-mineral and diluted nonbuffered solutions. Acidic peatlands harbor a large diversity of methanogenic archaea. Despite the ubiquity of methanogens in these peatlands, indigenous methanogens capable of growth at acidic pH values are resistant to culture. Only recently, methanogens belonging to an uncultured family-level clade in the Methanomicrobiales and to genus Methanobacterium have been isolated.

1

Introduction

The soil–microbe system is one of the most diverse components of the terrestrial ecosystem. Soil is an extremely complex, variable living medium performing many vital functions such as biomass production and storage and microbial transformation of various substances, including water and carbon. These processes are of particular interest in freshwater wetland ecosystems where nutrient cycling is highly responsive to fluctuating hydrology, and gases produced in soil may contribute to climate warming. Northern peatlands are one of the most typical environments located in Eurasia and North America. This type of environments has been proved to be one of the powerful sources of atmospheric methane (Matthews and Fung, 1987). Besides low pH, other extreme conditions of such environments are high content of organic matter, low mineralization, and permanently low temperature in anoxic peat layers. Methane emission from such environments has been well established by various field measurements (Avery et al., 1999; Bellisario et al., 1999; Williams and Crawford, 1984). The ecological, biogeochemical, and hydrological regimes in peatlands are complex, resulting in a wide temporal and spatial variability of CH4 emissions. The rates of methanogenic degradation processes go down significantly under acidic conditions. Relatively, little is known about the relationship between mechanisms of methane production and methanogenic community composition. In anaerobic peat, CH4 production is considered to be dominated by acetoclastic and H2-dependent pathways. Until very recently, there were only limited data about methanogenic anaerobic community developing in wetlands, particularly in acidic peatlands.

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Acidophilic microorganisms inhabiting peat wetlands can have peculiarities in the regulation of metabolism as well as in specific physiological adaptations to low pH (Russel, 1991). Molecular biological studies of wetland anaerobic communities have revealed physiologically distinct bacterial and archaeal lineages also (Bra¨uer et al., 2006a; Dedysh et al., 2000; Galand et al., 2002; Utsumi et al., 2003).

2

Anoxic Waterlogged Soils as a Habitat for the Methanogenic Community

Soil is defined as the top layer of the earth’s crust, capable of supporting life. It is the most complicated biomaterial on the planet and is formed by mineral particles, water, air, decaying organic matter (humus), and living organisms (Crawford et al., 2005). Soil is the greatest reservoir of biodiversity on the planet. Prokaryotes comprise more than half of the biodiversity on Earth, and their diversity in soil has been estimated to be about three ordersofmagnitude greater than in all other environments combined (Curtis et al., 2002). Soil microorganisms mediate many processes, providing the turnover of elements such as carbon, nitrogen, sulfur, and different metals that regulate ecosystem function and also feed back to influence atmospheric chemistry. The waterlogging of biologically productive soils makes them anoxic and allows anaerobic microbial community to develop. Once the soil becomes anoxic, remineralization rates lower and organic matter can accumulate. Under anaerobic conditions in freshwater environments, CH4 production becomes the most important terminal electron sink of anaerobic respiration. Wetlands are areas on which water covers the soil or where water is present either at or near the surface of that soil. Water can also be present within the root zone all year or just during various periods of the year. The result is a hydric soil, one characterized by an absence of free oxygen sometimes or always. Wetland ecosystems are characterized by hydrophilic plant communities and have fluctuating hydrology that gives rise to an interplay between aerobic and anaerobic processes (Gutknecht et al., 2006). Wetlands, because of their complex hydrology and nutrient cycling and presence in both urban and unmanaged areas, are uniquely positioned to influence biogeochemical cycling in many regions and at many scales. A wetland may be found in coasts, estuaries, floodplains, shallow lakes, and peatlands. Wetlands are considered to be the largest natural source of global atmospheric CH4 and are responsible for the release of ca. 10–30% (50–150 Tg CH4) of the total annual methane emission (Cicerone and Oremland, 1988; Matthews and Fung, 1987). Wetlands are not equally distributed across latitudinal zones, and therefore, wetlands will have a different impact on the CH4 budget at different latitudes. The greatest areal extent of wetlands is at higher latitudes of the Northern Hemisphere (>40 N) occupied by peatlands (Aselmann and Crutzen, 1989). Peatlands are unbalanced wetland ecosystems where productivity exceeds biodegradation. This imbalance leads to the accumulation of organic deposits (peats), which are derived from dead and decaying plant material under conditions of permanent water saturation. Although peatlands represent a relatively small area (3% of the Earth’s surface) and have low levels of primary productivity compared with other terrestrial ecosystems, the peataccumulating wetlands are significant repositories of carbon. They store more than 30% of the world’s terrestrial C pool and represent a large natural source of CH4 to the atmosphere (Matthews and Fung, 1987; Whalen, 1993).

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Emissions of CH4 from northern peatlands vary as a function of temperature, pH, substrate and nutrient availability, anoxia due to flooded conditions, the degree of CH4 oxidation that occurs in the upper aerobic sediments of peatlands, production enhancement and transportation via certain vascular vegetation (Bellisario et al., 1999; Whalen, 2005), and the presence of other microorganisms outcompeting the methanogens such as sulfate and iron-reducers and homoacetogens (Achtnich et al., 1995; Frenzel et al., 1999; Kotsyurbenko et al., 1996, 2001). There are three primary mechanisms for CH4 and CO2 transport from anoxic soils to the atmosphere: diffusion, ebullition, and movement through plant tissues (Lansdown et al., 1992). Plants may enhance the emission of CH4 through root-leaf transport and bypassing oxidation in the aerobic zone as well as by releasing carbon exudates from plant roots, which are labile substrates for methanogens (Stro¨m et al., 2003). Peatlands include a wide range of ecosystems: each with a characteristic peat soil derived from partially decaying plant material and with little or no rock-derived minerals. Types of peatlands mostly depend on geographic region, terrain, and vegetation type. A characteristic of many peatlands is that the plant species composition of surface vegetation can be quite different from the remains of plants in the peat that dominated in the past. Deep peat deposits occur in wetlands in northern latitudes and the depth of the peat decreases with a decreasing latitude due, in part, to climate (Yavitt et al., 1987). Peatlands are characterized by surface heterogeneity and fluctuating water table position that results in differences in the thermal regime, nutrient cycling, plant community composition, and organic matter production on a scale of several meters (Whalen and Reeburgh, 2000). A major distinction in the types of peatlands is between bogs and fens (Aselmann and Crutzen, 1989). Bogs are the most acidic peatlands, which are fed only by precipitation and are nutrient-poor. They are characterized by low mineral nutrient concentrations and dominated by Sphagnum mosses and a few ericaceous shrub species. Fens are near-neutral-pH peatlands, which are fed by surface and ground water as well as precipitation and tend to be more nutrient-rich. An increasing pH is associated with less Sphagnum and a dominance by Carex sedges and graminoid plants. Except for the surface water microlayer, wetlands are anaerobic environments. As such, they represent suitable habitats for all microbial groups from the methanogenic community (Conrad, 1996). Sulfate reduction zone can be also extensive in wetland soils, especially in marshes that are influenced by sea-water. Methane production may be limited by microbial iron reduction (Metje and Frenzel, 2005). Fe(III)-reducing microorganisms may suppress methanogenesis competing for H2 or acetate (Roden and Wetzel, 2002). However, most wetland soil is devoid of O2 and also contains no electron acceptors other than CO2 and H+.

3

Methanogenesis in Peatlands

The activity of methanogens in peatlands can easily be detected, because methanogenic metabolic activity is directly related to the amount of CH4 produced in peat. Measurements of potential methane production show that methanogenic activity is restricted to the waterlogged layers of the peat. The maximum CH4 production in peat profile takes place at the depth where most of the anaerobic degradation occurs.

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The methane production in acidic peats has been shown to be stimulated by increasing temperature (Kotsyurbenko et al., 2004) and pH (Kotsyurbenko et al., 2007; Williams and Crawford, 1984). The effect of these environmental parameters on methanogenesis indicates that peatland methanogens are metabolizing under suboptimal growth conditions that indicate the important role of microbial adaptation. The fact that microorganisms have growth optima that can never be achieved under in situ conditions is well established for different ecosystems. However, it has been also shown that peatlands contain endemic acid-tolerant microorganisms participating in methane cycle at a low pH (Dedysh et al., 2000, 2002). Methane is a major product of the degradation of organic matter in peatlands. Methane is almost exclusively produced from either acetate or from H2/CO2 (Conrad, 1999). Methanol, which is released during the decomposition of pectin, a polymer of methoxylated galactouronic acid and a major cell wall component of plants, usually plays only a minor role (Hines and Duddleston, 2001). The relative contribution of H2/CO2 versus acetate as methanogenic precursors can be quite different in various wetlands. The predominance of one respiratory pathway over another is generally a result of either the availability or lability of the carbon substrate. The acetate fermentation pathway is thought to dominate over CO2 reduction when fresh organic material is utilized as in sites with high plant productivity. Less productive plant communities with more recalcitrant material (Sphagnum dominate, sedges are scarce) tend to use the CO2 reduction pathway (Galand et al., 2005; Keller and Bridgham, 2007). Aceticlastic methanogenesis seems to predominate in fens populated by Carex sedges (due to the availability of root exudates supplied by the vascular plant community), while CO2 reduction was more important in Sphagnumdominated bogs (Kelley et al., 1992). The acetate fermentation often exceeds CO2 reduction in summer when decomposition of organic matter is most active (Avery et al., 1999). Enriched d13C-CH4 isotopic signatures attributable to methane production pathway via acetate (Whiticar, 1999) are often associated with sites exhibiting high rates of plant production and large CH4 fluxes. This confirms that abundant fresh organic material at sites with the greatest plant productivity stimulates larger CH4 emissions via the acetate fermentation pathway. The methanogenic pathways also have been shown to be dependent on pH and temperature. A low pH is advantageous to H2-dependent methanogenesis, whereas a low temperature is favorable to acetoclastic methane pathway (Conrad, 2002; Kotsyurbenko et al., 1996, 2005, 2007). Acetate can even accumulate seasonally in northern peatlands (Duddleston et al., 2002; Hines and Duddleston, 2001), followed by oxidation, and it has been proposed that aceticlastic methanogenesis may be absent or inhibited at these sites. Acetate also can be oxidized to CO2 via aerobic respiration or other oxidative microbial processes (e.g., via the dissimilation of iron or nitrate) or oxidized syntrophically to CO2 by the concerted activity of acetate-oxidizing anaerobes (Nu¨sslein et al., 2001). Supplemental acetate can be inhibitory to methanogenesis in acidic peat samples, whereas glucose and H2 can be stimulatory (Williams and Crawford, 1984). The negative influence of humic substances on methanogenesis has also been reported (Stewart and Wetzel, 1982). Acetic acid and other fatty acids are known to be toxic at a low pH (Russell, 1991). Acetate and other volatile fatty acids inhibit methanogenesis in bog peat at pH 4.5, but not at pH 6.5 (Horn et al., 2003). The explanation is the abundance of undissociated acetic acid under low pH conditions. At pH below 6.0, a greater fraction of total acetate will be present as acetic acid (pKa = 4.7), which can permeate cell membranes, causing acidification of the cell interior and

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acts as a decoupler of the proton motive force that can be lethal to the cell (Russell, 1991). Acetate concentrations of 5–10 mM and higher are considered to be inhibitory at a low pH, while acetate may be utilized at its natural concentrations in the micromolar range (Bra¨uer et al., 2004).

4

Methanogenic Diversity in Peatlands

The application of molecular techniques based on 16S rRNA and mcrA (encoding a subunit of the methylreductase involved in methanogenesis) gene sequences to study wetlands has revealed a diversity of methanogens belonging to the Methanomicrobiaceae, Methanobacteriaceae, Methancoccaceae, Methanosarcinaceae, and Methanosaetaceae as well as new archaeal lineages within the Euryarchaeota (Basiliko et al., 2003; Galand et al., 2002, 2005; Horn et al., 2003; Kotsyurbenko et al., 2004; Metje and Frenzel, 2007; Upton et al., 2000; Yavitt et al., 2006). The described representatives from these families are known to use both acetate and H2/CO2. It indicates that methanogenic community in peatlands contains all trophic groups of methanogenic archaea that are required to explain the formation of CH4 via acetoclastic and H2-dependent pathways. Nevertheless, methanogenic communities in bogs and fens are functionally different (Galand et al., 2002, 2005; McDonald et al., 1999). A bog has a more pronounced dominance of a few taxa, whereas a fen has a more even distribution among taxa. The methanogenic diversity at the bog is usually quite low (Galand et al., 2005). The peat pH, which is much lower in bogs, may be a factor in the selection of specific acid-tolerant microorganisms. Methanogenic community composition has also been found to change vertically within a site (Galand et al., 2002, 2005). The low concentration of acetate in mesotrophic peats favors Methanosaeta spp., which have a lower threshold for acetate than other acetotrophs belonging to the family Methanosarcinaceae (Galand et al., 2005). In ecosystems where acetate concentrations are high, Methanosaeta spp., are outcompeted by Methanosarcina spp. (Fey and Conrad, 2000). Essentially, a seasonal supply of fresh substrates in peatlands stimulates the growth of bacteria much more than the growth of archaea (Chan et al., 2005). The bacterial metabolism in turn stimulates methanogenesis, indicating the hydrolysis of polysaccharides and other complex organic materials as a first and rate-limiting step for the methanogenic degradation of organic matter production. Temperature also plays a role in the archaeal composition (Fey and Conrad, 2000). Methanogenic diversity appears to increase with an increasing temperature. The diversity of archaea among wetlands distributed across latitudinal gradients tends to increase with a decreasing latitude that also correlates with temperature (Utsumi et al., 2003). Thus, the succession of methanogenic archaea communities is dependent on the changes in environmental conditions and the quantity and quality of substrate, which are in turn determined by the plant community structure. Microorganisms inhabiting Sphagnum peat have specific growth requirements. They appear to be sensitive to mineral composition in the environment and can only develop at low ionic strength (Sizova et al., 2003). The sensitivity of the methanogenic population from wetlands to nitrates and sulfates has been reported (Scheid et al., 2003). Progress was recently achieved when acidophilic methanotrophs were isolated for the first time (Dedysh et al., 2000, 2002). Contrary to all previously known neutrophilic methanotrophs, they have maximal growth at pH 4–5 and are very sensitive to nitrates (Dedysh, 2002).

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Acidotolerant Methanogens

Attempts to isolate acidophilic or acidotolerant methanogens typical of a wetland ecosystem in pure culture have failed until very recently, although acid-tolerant enrichment cultures have been reported (Bra¨uer et al., 2006b; Horn et al., 2003; Sizova et al., 2003; Williams and Crawford, 1985). Nevertheless, the in situ emission of CH4 from the bogs at pH < 5.0 indicates that methanogens are functional under these acidic conditions. Additionally, methanogenic archaea rely on other organisms, usually bacteria, to convert the organics to methanogenic precursors. They often grow in association with other microorganisms and can be hardly separated in culture. Moreover, bacterial satellites probably provide adequate redox potential by the removal of traces of oxidants, and probably, supply growth factors other than regular vitamins (Sizova et al., 2003). Hence, the major methodological principle should be to mimic the natural environment of methanogens by developing media that more closely approximate the in situ conditions (Sizova et al., 2003). Recent progress has been made in the successful isolation of moderately acidophilic methanogens belonging to Methanomicrobiaceae and Methanobacteriaceae families (Bra¨uer et al., 2006a; Kotsyurbenko et al., 2007). The former methanogen is a member of an uncultured family-level clade that is prevalent in many acidic peat bogs in the Northern Hemisphere. Both isolates have the optimum pH for methanogenesis near 5.0, which is lower than that of any previously described methanogen. The methanogens are able to metabolize at pH 4.0 and at 10 C. It points out their capacity of developing under in situ conditions and their active participation in methane cycle in wetlands. They are hydrogenotrophic methanogens that confirm the importance of this methanogenic pathway for acidic wetlands.

6

Research Needs

Current information on methanogenesis in wetlands is insufficient for making adequate and precise predictions of greenhouse gas emissions on the global level. A detailed knowledge of the structure and the functioning of methanogenic communities from the largest terrestrial methane-producing ecosystems would be beneficial for understanding the microbial ecology of methane production, which, in turn, may lead to interventions to ultimately control the factors of the global CH4 turnover. The research is necessary that aims at integrating the analysis of environmental factors, biogeochemical processes, and bacterial and archaeal community profiles in diverse wetland ecosystems. The effective concept of such study includes a consecutive consideration of interacting biosystems of different complexity levels (the system approach). Accordingly, methane fluxes measurements in situ and in peat bog samples (ecosystem level) are followed by experiments on trophic microbial interaction, predominant methanogenic pathways (community level), and finally, characterizing the microbial diversity and the metabolic potential in partial methanogens in the community, (microbial group level) with an attempt to investigate the key microorganisms (microorganism level). Both cultivation-dependent and -independent approaches should be involved to characterize the true in situ microbial composition. The specific requirements of different groups of microorganisms living in mineral-deficient wetlands at low temperatures and an acidic pH should be analyzed to establish new protocols for getting active enrichments and maintenance of important, yet poorly culturable, microorganisms, which will significantly

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expand our knowledge on biodiversity and the limits to life under the harsh environmental conditions. Microorganisms developing under such extreme conditions may exhibit novel adaptation mechanisms, which could lead to new bioproducts and extend the range of biotechnological applications.

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51 Environmental Constraints that Limit Methanogenesis T. Hoehler1 . R. P. Gunsalus2 . M. J. McInerney3,* 1 Exobiology Branch, NASA Ames Research Center, CA, USA [email protected] 2 Department of Microbiology and Molecular Genetics, University of California Los Angeles, Los Angeles, CA, USA [email protected] 3 Department of Botany and Microbiology, University of Oklahoma, Norman, USA *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 636

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Biochemical and Bioenergetic Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 639

3 Ecological Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641 3.1 Syntrophic Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 641 3.2 Competitive Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 642 4 4.1 4.2 4.3 4.4 4.5

Physicochemical Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 645 Oxygen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 645 Temperature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 645 pH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 647 Salinity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 648 Pressure . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 649

5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 650

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_51, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Methanogens are represented across a diverse range of aquatic ecosystems, including habitats that are driven principally by geochemically-supplied substrates (e.g., hot springs and hydrothermal vents, volcanically-influenced settings, and, potentially, the deep crustal subsurface) and those dependent on substrates derived from organic matter (e.g., aquatic sediments, wetlands, agricultural or natural soils subject to inundation, sewage digesters, and the anoxic portions of animal digestive tracts). Similarly, they are tolerant to a broad range of physicochemical conditions, including temperatures from 0 to 122
C, pH values of 3–10, salinities from 0 to halite saturation, and pressures of at least 75 MPa. In general, methanogen distribution is constrained by ecological interactions (both stimulatory and competitive) and/ or physicochemical environmental factors that act at biochemical or bioenergetic levels. In addition to physicochemical ‘‘extremes’’ (principally temperature, salinity, and pH), the environmental distribution of methanogens is constrained to a large degree by energy availability and by environmental distributions of oxygen (biochemical inhibition) and the seawater anion sulfate (competitive effects that act at a bioenergetic level). Although methanogen tolerances to individual extremes are documented in culture, and the corresponding biochemical adaptations understood to varying degrees, the natural environment frequently presents combinations of extremes, combined with energy limitation, that may serve to limit methanogen distribution to less than the optimally tolerated range. Little is understood about the compound effects of such extremes, nor the commonalities among them that will ultimately form the basis for predictive models of environmental population distribution. Future work that targets these questions, through a combination of culture work, in situ studies, and theoretical (conceptual and quantitative) models, represents the way forward in better understanding the physiological ecology of methanogens.

1

Introduction

Biological production of methane, also termed methanogenesis, is a quantitatively important component of the global carbon cycle on the modern Earth (Hedderich and Whitman, 2006), and has likely been so since throughout most of the history of Earth’s biosphere. Averaged annually, about 1–2% of present global net photosynthetic carbon fixation is ultimately processed through methanogenesis (Hedderich and Whitman, 2006). Of the 500–600 Tg of methane that enter the atmosphere annually (Ehhalt et al., 2001), over 70% (350–400 Tg), derives from microbial activity. For at least several decades, atmospheric methane concentrations have been increasing at a rate of 1–2% annually (Reeburgh, 2003), owing principally to anthropogenic effects (agriculture, biomass burning, land use changes, and ecosystem effects attributable to global climate change). Although the abundance of methane in the modern atmosphere is less than one percent of the abundance of CO2, it is considerably more efficient, on a per-molecule basis, as a ‘‘greenhouse’’ gas (Ramanathan et al., 1985). Because of this important and continually increasing role in Earth’s radiation budget, the global methane budget and its associated microbial cycles have received considerable study in recent decades. Importantly, from the perspective considered in this chapter, much of this work has sought to understand the impact of changing physicochemical environment on methane production and emission. In particular, a variety of studies have focused on understanding the influence of global change-related effects in areas that stand to be significantly impacted (e.g., melting of permafrost, seawater flooding of wetlands, and land use change).

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51

Methane may also have played an important role in the global carbon cycle and associated climate dynamics throughout much of Earth’s history. Microbial methanogenesis is thought to have arisen as early as 3.5+ billion years (Ueno et al., 2006). Proportionately, it’s role in carbon cycling may have been much more important on the early versus the modern Earth (Catling et al., 2001), owing to the relative absence of oxygen and sulfate, two of the principal limiters of the methanogenic biosphere on the modern Earth (as discussed in subsequent sections of this chapter). Indeed, methane (and underlying methanogenic activity) is hypothesized to have played a dominating role in Earth’s early climate dynamics, potentially warming the planet during a period in which global glaciations may otherwise have occurred (Kasting et al., 2001; Pavlov et al., 2003). Significant and relatively abrupt changes in atmospheric methane during Earth’s history have similarly been implicated in global climate change (e.g., ‘‘snowball Earth’’ events associated with rapid depletion of atmospheric methane or significant warming events associated with large scale release of hydrate methane to the atmosphere). Studies of historical methane-driven climate dynamics focus largely, as with present day studies, on the effects of changing physicochemical environment and associated ecosystem dynamics on methane production and cycling. The methanogen biosphere occupies a diverse array of ecosystem types (Wolfe, 1971) and a broad range of physicochemical conditions. The best known and (at the Earth’s surface) most pervasive methanogenic niches are in oxygen-free aqueous systems that provide a supply of organic matter (the ultimate source, via microbially-mediated decomposition, of the substrates principally utilized by methanogens). Such systems include marine, lacustrine, and riverine/estuarine sediments; wetlands, such as swamps, bogs, and periodically flooded forest soils; agricultural soils subject to inundation, such as rice paddies; sewage digesters; and the anoxic portions of animal digestive tracts (see, e.g., Chaban et al., 2006; Hedderich and Whitman, 2006; Liu and Whitman, 2008). Deep sea sediments and oil reservoirs also contain active methanogenic communities (Jones et al., 2008; Rossel et al., 2008; Wilhelms et al., 2001). Additionally, because H2 – the substrate utilized by the broadest range of methanogens – is produced almost ubiquitously through the interaction of crustal rocks and water (see review in Hoehler, 2005), the potential exists for a pervasive methanogen biosphere that is supported not by photosynthetic productivity, but by geochemical energy sources. Methanogenic activity is documented at temperatures from 0 to 122
C, at pH from 3 to 10, at salinities from near 0 to >5 M NaCl, and at pressures greater than 75 MPa (> Table 1). The pervasiveness of methanogens across diverse ecosystems and, in particular, the tolerance to wide-ranging ‘‘extremes’’ may arise from a combination of factors, including: (1) the availability of methanogenic substrates in a wide variety of settings, including from geochemical sources; (2) a relatively simple biochemical machinery (where a smaller genome, fewer core enzymes, and less complexity in general may foster tolerance of a broader range of physicochemical conditions and/or more rapid adaptation to new conditions); and (3) more than three billion years in which to adapt and evolve a variety of phenotypes around a simple core metabolism. Together, these factors have presented methanogens with the impetus, potential, and time to explore and occupy, through evolution, an extremely broad range of ecological niches. Despite this ecological plasticity, the distribution of methanogens in nature is quite limited in comparison with the distribution/availability of potential methanogenic substrate. In general, methanogen distribution is constrained by ecological interactions or physicochemical environmental factors that breach biochemical or bioenergetic limits. This chapter considers these limitations and the resulting major environmental controls on methanogenesis. The interested reader is also directed to Zinder (1993) for a thorough consideration of the physiological ecology of methanogens.

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. Table 1 Examples of extreme methanogens and syntrophic metabolizers Extreme Hyperthermophilic methangens

Organism




Range







Reference

Methanopyrus kandleri

98 C; 105 C 84–110 C; at 40 MPa 90–122
C at 40 MPa

Kurr et al. (1991); Takai et al. (2008)

Methanothermus fervidus

83
C

65–97
C

Stetter et al. (1981)

Methanothermus sociabilis

88
C

55–97
C

Lauerer et al. (1986)

Methanocaldococcus jannaschii

85
C

50–91
C

Jones et al. (1983)

Methanocaldococcus infernus

85
C

55–91
C

Jeanthon et al. (1998)

Methanocaldococcus vulcanius

80
C

49–89
C

Jeanthon et al. (1999)

Methanocaldococcus fervens

85
C

48–92
C

Jeanthon et al. (1999)

Methanotorris igneus

88
C

45–91
C

Burggraf et al. (1990)

Extreme thermophilic Thermosyntropha syntrophic lipolytica metabolizers

Psychrophilic/ psychrotolerant methanogens

Optimum

60–66
C pH 52–70
C pH 7.5– Svetlitshnyi 8.1–8.9 9.5 et al. (1996)

Desulfotomaculum thermocisternum

62
C

41–75
C

Nilsen et al. (1996)

Pelotomaculum thermopropionicum

55
C

37–70
C

Imachi et al. (2002)

Syntrophothermus lipocalidus

55
C

45–60
C

Sekiguchi et al. (2000)

Methanogenium frigidum

15
C

0–18
C

Fransmann et al. (1997)

Methanococcoides burtonii

23
C

1.7–30
C

Fransmann et al. (1992)

Methanosarcina lacustris

25
C

1–35
C

Simankova et al. (2001)

Methanosarcina baltica

25
C

4–27
C

von Klein et al. (2002)

Methanosarcina acetivorans

34–30
C

~ 9
C

von Klein et al. (2002)

Methanocorpusculum strain MSP

25–35
C

1–35
C

Simankova et al. (2003)

Methanomethylovorans hollandica

25–35
C

1–35
C

Simankova et al. (2003)

Methanogenium marinum

25
C

5–25
C

Chong et al. (2002)

Environmental Constraints that Limit Methanogenesis

51

. Table 1 (Continued) Extreme

Organism

Optimum

Range

Reference

Psychrophilic/ psychrotolerant syntrophic metabolizers

Algorimarina butyrica

15
C

10–25
C

Kendall et al. (2006)

Acid tolerant methanogens

‘‘Candidatus Methanoregula boonei’’

pH 5

pH 4.3–5.8

Bra¨uer et al. (2006)

‘‘Candidatus Methanosphaerula palustris’’

pH 5.3–5.5

pH 4.8–6.5

Cadillo-Quiroz et al. (2008)

Methanobacterium espanolae

pH 5.5–6.0

pH 4.7

Patel et al. (1990)

Methanohalophilus zhilinae

pH 9.2

pH 8.0–10

0.7 M NaCl

0.2–2.1 M NaCl

Mathrani et al. (1988)

Methanolobus oregonensis

pH 8.6

pH 8.2–9.2

Liu et al. (1990)

Methanolobus taylorii

pH 8

pH 5.5–9.2

Oremland and Boone (1994)

Methanohalobium evestigatum

4.3 M NaCl

2.6–5.1 M NaCl

Zhilina and Zavarzin (1987)

Methanohalophilus mahii

1.0–2.5 M NaCl

0.5–3.5 M NaCl

Paterek and Smith (1988)

Methanohalophilus halophilus

1.2–1.5 M NaCl

0.3–2.6 M NaCl

Zhilina (1983)

Halomethanococcus doii

3.0 M NaCl

>1.8 M NaCl

Yu and Kawamura (1987)

Methanohalophilus portucalensis

0.6–2.1 M NaCl

>1.4 M NaCl

Boone et al. (1993)

Methanocaldococcus jannaschii

75 MPa

Miller et al. (1998)

Methanococcus thermolithotrophicus

50 MPa

Bernhardt et al. (1988)

Akalophiles/alkaline tolerant

Halophiles/ halotolerant

Piezophiles

2

Biochemical and Bioenergetic Considerations

Biochemical limitations are encountered principally through physical or chemical disruption of core metabolic molecules, structures, networks, or processes. Examples include the thermal destabilization of enzyme tertiary structure, enhanced chemical hydrolysis of biopolymer linkages, chemical inactivation of enzyme binding sites, or the inherent limitation of enzymes to take up and process substrates at levels needed to compete effectively or support metabolism. Specific biochemical effects on methanogens are considered below. Bioenergetic constraints on environmental habitability arise from life’s fundamental need to harness energy from the surroundings and invest it in order to maintain what is, ultimately,

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a disequilibrium state (in the form of complex biological molecules and structures that are thermodynamically unstable with respect to the general environment). For an environment to be habitable from an energetic standpoint, the provision of energy by that environment and the organism’s capability to access and use that energy must balance or exceed the organism’s demand for energy (Hoehler, 2004, 2007; Shock and Holland, 2007). The biological demand for energy is manifest in two requirements, which are analogous to the voltage and power requirements of an electrical device. The analog to voltage (energy per unit of energy carrier) is the biological energy quantum (BEQ), which is defined as the smallest free energy change of a metabolic reaction that can still be used to drive ATP synthesis (upon which sustained metabolic activity depends) (Schink, 1997; Schink and Stams, 2006). The BEQ requirement is described more thoroughly, and with specific reference to methanogens, in the introductory chapter on microbial hydrocarbon production and bioenergetics. The analog to power (energy per unit time) is the maintenance energy (ME), which defines the flux of energy needed to support a unit of biomass in a steady state at a net zero growth rate (Harder, 1997; Tijhuis et al., 1993). Each requirement is characterized by a minimum value, below which sustained metabolism is not possible and both requirements must be met in order for a given environment to be habitable with respect to specific organism (Hoehler, 2004, 2007) (> Fig. 1). The magnitudes of the BEQ and, in particular, ME requirements are set in part by the biochemical and physiological characteristics of the organism in question and in part by the environment in which it lives. Nominally, the magnitude of the BEQ has been estimated at about 20 kJ·mol1 for actively growing organisms (Schink, 1997) and 12 to 15 kJ·mol1 for organisms operating under energy-limiting conditions (Schink and Stams, 2006). Measurements of energy yields associated with methanogenesis in various environments suggest that methanogens may be able to capitalize on energy yields of about 10 kJ·mol1 (see summary in Hoehler, 2004). The magnitude of the ME requirement may vary more substantially across different organisms, and is considerably less well constrained than that of the BEQ. Estimates of ME derived from culture-based and environmental studies, and from growing versus non-growing organisms vary over orders of magnitude (Morita, 2000; Price and Sowers, 2004; Tijhuis et al., 1993). For both types of energy requirement, but particularly for the ME requirement, environmental deviations from biologically optimal physiochemical conditions – the classical ‘‘extremes’’ of temperature, pH, salinity, radiation, and so forth – may increase an organism’s energy demands significantly. Indeed, natural systems can present multiple physicochemical extremes (e.g., high temperature and low pH) that may have compound effects in increasing cellular energy demands. For environments offering only limited energy supplies (in the form of limited fluxes of methanogenic substrates), these increasing energy demands may ultimately exceed the environmental energy supply, and thereby render the environment uninhabitable (Hoehler, 2007). Thus, in natural systems, bioenergetic factors may limit habitability before absolute biochemical limitations – which are typically determined in cultures under ideal, energy-rich conditions – are encountered. While the general bioenergetic considerations outlined above are applicable for all organisms, they are especially relevant for metabolisms offering relatively low energy yields under typical environmental conditions. Methanogenesis exemplifies this situation, and many aspects of the environmental distribution/limitation on methanogen activity are attributable to bioenergetic effects.

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51

. Figure 1 The importance of energy yield and energy flux in determining the habitability of an environment (after Hoehler, 2007). The free energy of the catabolic reaction (DGrxn) must be more favorable (e.g., more negative DG) than the minimum amount of energy needed to make ATP (biological energy quantum, BEQ). The flux of energy must be larger than the minimum amount to maintain cellular functions and viability (maintenance energy minimum, MEmin). Growth will occur when the energy flux is large enough to supply sufficient energy for biosynthesis (darker shaded region). Habitable environments (lightly shaded region) are those where that catabolic reaction is more favorable that the BEQ and where energy flux is sufficient to maintain viability but insufficient for growth. If either requirement is not met by the energy available in the local environment, the system is uninhabitable (unshaded region). Note that the magnitude of both BEQ and particularly maintenance energy are highly dependent on physicochemical environment and may, in the case of environmental extremes, rapidly exceed levels that can be sustained within a given system.

3

Ecological Interactions

Methanogens as an overall metabolic group are capable of using H2/CO2, formate, acetate, CO, and several methlyated compounds as substrates for methane production, although individual taxa may use only a subset (Zinder, 1993). While any of these may conceivably predominate in a given environment, the overall most quantitatively important methanogenic substrates are H2/CO2 (and/or formate) and acetate. Both occupy a central role in carbon and electron flow in anaerobic microbiology, and in the metabolic processes of a variety of microorganisms. As such, they present a basis for interactions, both stimulatory and inhibitory, between methanogens and other organisms.

3.1

Syntrophic Interactions

Methanogens’ ability to grow autotrophically using H2 as an electron donor potentially allows them to directly access geochemical sources of energy (e.g., the H2 produced by water-rock

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reactions). However, most of the known methanogenic ecosystems (including marine, lacustrine, and riverine/estuarine sediments; wetlands, such as swamps, bogs, periodically flooded forest soils; periodically flooded agricultural lands, such as rice paddies; sewage digesters; and the anoxic portions of animal digestive tracts; see, e.g., Chaban et al., 2006; Hedderich and Whitman, 2006; Liu and Whitman, 2008) are instead fueled by decomposition of complex organic matter. In the absence of oxygen, the complete decomposition of complex organics requires the collective activities of a diversity of anaerobic microorganisms, each catalyzing individual steps in the overall process (Schink, 1988). Only in the final steps of this process are methanogenic substrates (principally acetate and H2) generated, so that methanogen activity in such environments is intimately linked with and dependent on the activity of syntrophic partner organisms. This partnership is particularly critical in reference to the exchange of H2 between partner organisms. This process, termed ‘‘interspecies hydrogen transfer’’ exerts significant control on the pathways of carbon and electron flow in organic matter degradation, by virtue of the strong influence H2 exerts on the thermodynamics of microbial H2-cycling metabolisms (e.g., Bryant et al., 1967; Iannotti et al., 1973; Wolin and Miller, 1982). (The thermodynamic considerations associated with interspecies electron transfer are considered in greater detail in the introductory chapter on microbial hydrocarbon production and bioenergetics) Because of this dependence, any environmental or ecological factor that can influence the functional members of the overall organic matter-degrading community (in particular, the syntrophic partners) can also influence the conduct of methanogenesis. Thus, the environmental distribution of methanogenesis may be limited by community-level effects before the absolute physicochemical tolerances of methanogens are actually encountered. This is principally a bioenergetic limitation on methanogen distribution, by virtue of disruption of substrate flow to methanogens at levels or rates needed to meet cellular energy demands.

3.2

Competitive Interactions

Acetate and H2 are utilized in a wide range of microbial metabolisms, so that methanogen distribution may be limited by competition for these substrates. In anoxic systems driven by organic matter decomposition, the principal competitors for acetate and/or H2 are organisms that oxidize these substrates using a range of electron acceptors. In general, the most environmentally relevant of these are the inorganic electron acceptors nitrate, Mn4+, Fe3+, and sulfate (Nealson and Saffarini, 1994; Zehnder and Stumm, 1988), although a variety of other organic and inorganic oxidants can serve the same function and may be important in specific environmental settings. Observations of anoxic sediments show that organic matter remineralization often proceeds via successive oxidants, in the order indicated above, with one oxidant (e.g., sulfate) being completely exhausted before another one (e.g., CO2, in methanogenesis) is utilized. The ordering of oxidants above also reflects the magnitude of the standard Gibbs free energy yield associated with oxidation of hydrogen or acetate by that oxidant, with nitrate delivering the largest standard free energy yields and CO2 (to methane), the smallest (Zehnder and Stumm, 1988). Thus, according this general notion, the presence of any of the oxidants mentioned above would completely exclude methanogenesis. In practice, the presence or absence of sulfate, a major anion in seawater, is by far the most important among these oxidants in limiting the environmental distribution of methanogenesis as it effectively limits the distribution of methanogenesis in surface sediments of marine, estuarine, and hypersaline systems, which collectively constitute a large repository of potential methanogenic fuel.

Environmental Constraints that Limit Methanogenesis

51

As suggested by the ordering of oxidants by energy yield, the competitive exclusion of methanogenesis is hypothesized to have a thermodynamic/bioenergetic basis. Differences in standard free energy yields by themselves, however, do not provide a mechanism for exclusion of one organism by another. Rather, a larger free energy yield potentially enables one organism to compete more effectively for a common substrate, such as H2 or acetate, because it can (at least from a purely thermodynamic perspective) utilize the substrate to lower concentrations than organisms employing less energetic oxidants and still extract a free energy yield that meets the BEQ requirement. Complete inhibition of one metabolism by another will occur if one organism can actuate this potential and consume a common substrate down to concentrations that do not meet the BEQ or thermodynamic favorability requirements of a second. Complete inhibition of H2-consuming methanogenesis by H2-consuming sulfate reduction via this mechanism has been hypothesized or demonstrated in a variety of systems (e.g., CordRuwisch et al., 1988; Hoehler et al., 1998; Lovley and Goodwin, 1988), as exemplified in > Fig. 2. Whether or not the same potential is actuated in other microbial interactions (i.e., using acetate or other oxidants) depends on the energetically-advantaged organism

. Figure 2 Thermodynamic-based competitive exclusion of methanogenesis by sulfate reduction in a marine sediment (Cape Lookout Bight, North Carolina, USA). (a) Depth profiles of concentrations of sulfate (filled circles) and hydrogen (open circles). Note that hydrogen concentrations are maintained, by the activity of sulfate reducers, at five to ten fold lower levels within the sulfatecontaining zone. (b) Depth profiles of Gibbs free energies of reaction for H2-based sulfate reduction (per mole sulfate; filled circles) and methanogenesis (per mole methane; open circles). By virtue of their control over H2 concentrations, sulfate reducers limit methanogenic energy yields to values below the minimum bioenergetic requirement and, for the upper thirteen centimeters of the sediment column, at thermodynamically unfavorable levels (right of the solid vertical line at DG = 0). Below the depth of sulfate depletion (dashed horizontal line), methanogenesis yields about 10 kJ·mol1, consistent with lower-end estimates of the BEQ. In both (a) and (b), error bars represent the standard deviation about the mean of triplicate samples. (Figure modified from Hoehler et al., 2001.)

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having (1) enzyme kinetics that allow it to exercise its advantage (i.e., having enzymes that can take up substrates at the concentrations and rates required to exclude competitors) and (2) a supply rate of oxidant that exceeds that of H2 or acetate. Criterion (1) may limit the potential for competitive exclusion in the case of acetate, because the change in acetate concentrations required to actuate an energetic advantage is very large, and would require enzymes with capabilities to take up substrate at extremely low levels. For example, to actuate the >15 kJ·mol1 advantage that sulfate reducers appear to exercise over methanogens in the marine sediments presented in > Fig. 2 would only require a >4.5-fold decrease in H2 concentrations (as observed; Hoehler et al., 2001), but a >430-fold decrease in acetate concentrations. In practice, acetate concentrations may be only a few-fold lower, or less, in sulfate-reducing versus methanogenic sediments (Albert et al., 1991); hence, it is not clear that the same bioenergetic basis for competitive exclusion exists for acetate as in the case of H2. Nonetheless, methanogenesis (including from acetate) is often completely excluded by sulfate reduction. These seemingly discrepant observations can be reconciled if methanogenic metabolism of acetate is influenced by H2 concentrations in the system (and therefore by the actuated energetic advantage of sulfate reducers). Indeed, Finke et al. (2007) showed that methylotrophic methanogens can divert >95% of substrate methyl carbon (including from substrates not utilized by sulfate reducers) to CO2 and H2 (rather than to methane) when porewater H2 concentrations are held at low levels by sulfate reducers. This hypothesis could explain the apparent competitive exclusion of acetate-based methane production from sulfatecontaining sediments, despite the superficial lack of a thermodynamic basis, while also associating a potentially energy-yielding metabolic activity (Finke et al., 2007) with methylotrophic methanogens that appear to be present in some such sediments (Kendall et al., 2007). Criterion (2) may serve to limit the effectiveness of electron donors other than sulfate in excluding methanogenesis by competition for common substrates. For the dissolved electron acceptor nitrate, the principal concern is the total available pore water pool and the rate of replenishment. In many natural systems, nitrate is in short supply and is rapidly utilized (in either assimilatory or dissimilatory metabolism), so that its potential as a competitor for organic carbon resources is limited. However, in systems high in nitrate and largely lacking in sulfate (e.g., fertilized agricultural soils or wetlands affected by agricultural run-off), nitrate may be an important agent for exclusion of methanogenesis. For Mn4+ and Fe3+, the principal concern is that these electron acceptors occupy a largely insoluble phase (as particulate oxides), and so may be in short supply if the organisms that utilize them depend on diffusive supply of the dissolved form. As a result, the capability of Mn4+- and Fe3+-respiring organisms to lower the concentrations of common substrates in accordance with their energetic advantage may be limited. Lovley and Goodwin (1988) and Achtnich et al. (1995) observed decreased H2 concentrations in the presence of these metal oxides, but this effect may be highly concentration dependent (Hoehler, 1998). Indeed, metal reduction in sediments is frequently accompanied by co-occurring sulfate reduction or methanogenesis, except at high metal oxide concentrations (Thamdrup, 2000). Although bioenergetic considerations appear to underlie much of the competitive exclusion of methanogenesis (as dominated by the competitive effects of sulfate reduction), other mechanisms may also be important (Conrad et al., 2000; Lupton and Zeikus, 1984; Robinson and Tiedje, 1984). In particular, it has been suggested that low temperatures may favor acetogenesis (reaction of CO2/H2 to acetate, rather than to methane) over hydrogentrophic methanogenesis in some systems (Conrad, 1999; Kotsyurbenko, 2005; Nozhevnikova et al., 2007), even though methanogenesis is the more thermodynamically favored of the two

Environmental Constraints that Limit Methanogenesis

51

processes under most environmentally relevant conditions. It is suggested that homoacetogens may outcompete methanogens in this situation on the basis of better kinetic adaptation to low temperature, rather than on a bioenergetic basis (Kotsyurbenko et al., 2001).

4

Physicochemical Environment

Physicochemical ‘‘extremes’’ can impact methanogens at both biochemical and bioenergetic levels, either directly or by affecting their ecological interactions with other organisms.

4.1

Oxygen

The ambient concentration of oxygen is an important determinant of the environmental distribution of methanogenesis. Methanogens are strict anaerobes and, in culture, will not grow or metabolize (produce methane) in the presence of even trace levels of oxygen (Zinder, 1993). Given the high abundance of oxygen in Earth’s atmosphere, this sensitivity has the potential to severely limit methanogenesis. While this is certainly true in the sense that a much larger quantity of global primary productivity is processed by aerobic respiration than by methanogenesis, two factors serve to mitigate oxygen inhibition of methanogenesis to some degree. First, while methanogens do not grow or metabolize in the presence of oxygen, they do exhibit some tolerance to oxygen exposure at environmentally meaningful levels (Zinder, 1993). This suggests that core methanogen enzymes may not be irreversibly damaged or, at least, can be reactivated, following oxygen exposure. Oxygen tolerance may allow methanogens to persist in environments that may fluctuate between oxic and anoxic conditions (e.g., periodically flooded/aerated soils or sediments). Second, aqueous habitats containing particulate organic matter tend to endure limited permeation by oxygen because (1) the low solubility of oxygen in water – especially in saline water – effectively reduces the potential mass flux of oxygen from an overlying gas phase to methanogens inhabiting the aqueous phase; and (2) organic-containing sediments provide a physical matrix that limits oxygen transport to molecular diffusion (a slow process over distances greater than a few millimeters) and a biological/chemical sink for oxygen, via the reaction with organic matter to yield CO2. For systems having a high organic load (and, therefore, an abundance of potential methanogenic fuel), the combination of factors in (1) and (2) restricts oxygen penetration to a narrow surface zone that may range from microns to a few centimeters.

4.2

Temperature

Methanogens are represented across most of the known biologically-tolerated range of temperature from 0 to 122
C. Psychrophily. Methanogens are found in a variety of low temperature habitats, such as high latitude wetlands (e.g., boreal fens, tundras (See > Chapter 52, Vol. 1, Part 8), and bogs) and marine and freshwater sediments underlying deep (cold) waters and in arctic and antarctic regions (Caviccholi, 2006). As a group, these habitats also present significant differences in pH, salinity, pressure, and energy availability. Temperature ranges for cell growth in psychrophilic and psychrotolerant methanogenes generally span 20–35
C (> Table 1), and the lower limit can extend below 0
C when cells possess means to suppress ice formation

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(Caviccholi, 2006). Cell doubling times for Methanococciodes burtonii and Methanogenium frigidum are considerably below that described for the extreme thermophiles, and may be in the range of 0.1–0.3 generations per day (Caviccholi, 2006). The vast proportion of Earth’s surface that is subjected to periodically or permanently low temperatures (including most or all of the deep ocean basins, high latitude lakes and ocean shelves, and high latitude wetlands and soils), along with the significant reserves of organic carbon that are sequestered there, represent a vast expanse of potential habitat for psychrophilic methanogens. Nonetheless, relatively little is understood about the environmental diversity of the psychrohilic and psychrotolerant methanogens, or the mechanisms that allow them to adapt to low temperature niches. A combination of environmental studies and genomic analyses of model organisms will provide new approaches to address these questions (Saunders et al., 2003). Thermophily. Thermophilic and hyperthermophilic methanogens are found in fluid outflows from marine and fresh water volcanic seeps, hot springs, thermal mud pools and solfataric fields (Huber et al., 2000, and described in further detail in > Chapter 54, ‘‘Thermophilic methanarchaea inhabiting hot ecosystems’’). These habitats are typically rich in H2 and minerals, low in organics, and may vary significantly in pH and salinity (from fresh to marine). The chemoautotrophic methanogens Methanotorris igneus (Methanococcus igneus) and Methanothermus fervidus were isolated from a shallow offshore submarine vent and a thermal terrestrial waterhole in the mountains of Iceland, respectively. Methanopyrus kandleri strain 116, currently the high temperature ‘‘record holder’’ among cultured methanogens, was isolated from the Kairei hydrothermal field in the Central Indian Ridge at 2,400 m (T). It has a growth optimum of 105
C at 40 MPa, is capable of growth at 122
C at 40 MPa (> Table 1). By virtue of the general enhancement of metabolic rate by increasing temperature, cell doubling times for these thermophilic methanogens can be less than one per h (Jeanthon et al., 1998; Takai et al., 2004). The mechanisms of high temperature limitation of methanogens can be biochemical, bioenergetic, and/or ecological. While some large organisms can maintain internal temperatures significantly above or below ambient, individual microbes (or microbes in small clusters) cannot. Biologically meaningful temperature gradients cannot be maintained at the scale of microbial cells, so the environmental temperature is, effectively, the intracellular temperature. Thus, temperature effects can act directly upon the biochemical machinery of the cell. The deleterious effects of high temperatures relate principally to the thermal destabilization of core biomolecules, with resulting impacts on functionality (for example, the disruption of tertiary structure necessary for enzyme function). The biochemical impacts of high temperature for organisms in general (and applicable to methanogens) are thoroughly reviewed by Jaenicke and Sterner (2002). Such effects may set the ultimate upper limits on methanogen growth and metabolism in systems (especially cultures) that provide energy in abundance and optimize other growth parameters. Temperature can also influence methanogen metabolism by other than biochemical mechanisms. As noted earlier, for systems driven principally by organic matter decomposition, methanogen activity is ultimately dependent on the collective function of a broader community of (organic matter-degrading) organisms, and is therefore subject to the physicochemical limitations of critical organisms within that population. Thus, elevated temperatures may limit methanogen distribution by inhibiting partner organisms before the biochemical temperature limits of methanogens are encountered. Consistent with this notion, the cultured methanogens representing the upper end of the tolerated temperature range are generally derived from environments in which substrates (principally H2) are provided by geochemical

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sources, rather than by community-enabled organic matter remineralization. The maximum growth temperature for a cultured syntrophic metabolizer so far is about 75
C (> Table 1). Lastly, temperature has a strong effect on cellular maintenance energy, and may thereby serve to limit methanogenic activity via bioenergetic inhibition. The effect of temperature on maintenance energy has been quantified experimentally (Tijhuis et al., 1993) and conforms to an Arrhenius-type relationship (Harder, 1997) (equation 1): ME ¼ A  e Ea=RT where A is a positive constant, Ea is the activation energy, R is the universal gas constant, and T is temperature in K. According to this relationship, the energy required to support a unit of biomass increases exponentially with temperature. Empirically determined values for Ea (Harder, 1997; Tijhuis et al., 1993) predict that maintenance energy increases more than three-thousand fold as temperature increases from 0 to 100
C. Thus, in environments offering limited substrate fluxes, increasing temperatures may rapidly lead to bioenergetic limitation of methanogen growth and maintenance before absolute biochemical limits are reached.

4.3

pH

Methanogenesis is documented or inferred in cultures or environmental settings comprising a pH range of about 3–10 (> Table 1). Acidophilic methanogens are common to certain marine and freshwater boreal fens, tundras, and bogs, where accumulation of plant tannins and organic acids can lower pH to values ranging from weakly acidic to 3.5 or less (Zinder, 1993). These habitats are frequently also characterized by low temperatures, and so potentially present compound ‘‘extremes’’ to microbial inhabitants. As a group, acid-loving methanogens are relatively little studied, however observations in culture and acidic environments have helped to establish tentative pH limits on methanogenic growth and metabolism. Methane formation has been observed in peat samples (which could reflect either growth or metabolic activity without growth) at pH values as low as 3, although higher values are needed for optimal rates of methanogenesis or for growth in culture (Williams and Crawford, 1984). Some methanogens are also capable of growth or metabolism under alkaline conditions. Most of the studied alkaliphilic methanogens (e.g., those in > Table 1) are associated with evaporitic basins such as Mono Lake, California or the Dead Sea, the Rift Valley Lakes of East Africa, or desert soda lakes or streams, although methanogenic activity is also inferred in alkaline seeps where serpentinization yields waters with elevated pH (Kelley et al., 2005). Cultured representatives are moderately alkaliphilic (> Table 1), with one organism (Methanohalophilus zhilinae) having a pH growth optimum of 9.2 and a growth limit of 10 (Mathrani et al., 1988). Alkaline environments may present compound extremes for methanogen activity. Alkalinity associated with evaporitic settings is frequently accompanied by concentrated salts, so that some alkaliphilic methanogens are also halophilic. M. zhilinae, for example, is capable of growth at salinities up to 2 M NaCl, or about 4 times seawater salinity (Mathrani et al., 1988). Alkalinity may also be associated with high temperatures, as in alkaline hot springs or, in particular, hydrothermal settings associated with serpentinizing host rocks. For example, some venting fluids at the Lost City hydrothermal field reach pH values of 10–11 at temperatures of 70
C (Kelley et al., 2005). Significant deviations from neutral pH have the potential to adversely affect cellular biochemistry at a variety of levels, so that only modest variations in intracellular pH can be

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tolerated. However, habitation of environments having higher or lower than biochemically tolerable pH values is nonetheless feasible, because the lipid bilayer membrane (an effective barrier to ionic species like H+, OH or CO32) makes it possible to maintain intracellular pH at more moderate levels than in the extracellular medium, through active regulation. The mechanisms associated with such regulation are understood biochemically (Krulwich, 1995, 2000; Krulwich et al., 1996) and it is clear that actuation of these strategies (e.g., active transport of protons) must increase cellular maintenance energies. The effect of pH on maintenance energy in methanogens has not been quantified directly. However, the minimal energetic cost of pH regulation is determined by the rate of proton pumping necessary to maintain the appropriate internal pH (which, in turn, depends on the leakiness of the membrane and uncoupling by weak acids and bases) and the energy required to pump a unit quantity of protons (Krulwich, 2000). Both of these can be expected to increase with increasing or decreasing extracellular pH (increasingly large intracellular-extracellular pH differential), so that maintenance energies should increase monotonically with the degree of deviation of the environmental pH from a biochemical optimum. Thus, bioenergetic effects may factor prominently in setting the practical environmental pH limits. A secondary effect that may significantly constrain the habitability of alkaline or acidic environments with respect to methanogenesis is the speciation of methanogenic substrates in response to pH. Specifically, the conversion of methanogenic substrates into predominantly ionic forms that cannot diffuse across the cell membrane will require either energy-expending active transport of substrates or increases in membrane permeability, which could increase pH leakage and require higher rates of proton pumping to compensate. Such effects are probably most important in limiting methanogenesis in alkaline environments, with the deprotonation of acetic and carbonic acid, but could also conceivably be important for the methanogenic consumption of methylamines in acidic environments.

4.4

Salinity

Methanogens are found in environments with salinities ranging from fresh to halite-saturated (>5 M NaCl). Halophilic and extremely halophilic methanogens are most commonly associated with hypersaline environments that include dead seas, solar salterns and halite crystallizing ponds, and alkali lakes, where sodium chloride and other salts (e.g., magnesium choride and sulfate) may be highly concentrated (Ollivier et al., 1994; Oren, 2002). Notably, the conditions responsible for generating hypersaline conditions may also enhance alkalinity, and derive from elevated temperatures, so that methanogenic inhabitants of hypersaline environments may face multiple extremes. Described halophilic methanogens include moderate halophiles such as Methanohalophius mahii, Methanohalophilus halophilus, and Methanosalsus zhilinae (Ollivier et al., 1994), and extreme halophiles, including Methanohalobium evestigatum (> Table 1). The former have optimal growth with 1–2.5 M NaCl, while the latter are able to grow in saturating brines (over 5 M). Although elevated intracellular salt concentrations would directly and significantly impact cellular biochemistry, and organisms are known that have adapted to such conditions (the ‘‘salt in’’ strategists, e.g., Halobacteria), all of the known methanogens are ‘‘salt out’’ strategists (Oren, 2001). That is, intracellular salt concentrations are held below environmental levels by

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virtue of the barrier presented to ionic species by the lipid bilayer membrane, and through active transport of salts across the membrane. To compensate for the resulting differential in osmotic pressure, methanogens produce and concentrate intracellular osmolites such as glycine betaine, b-glutamate, b-glutamine, and Ne-acetyl-b-lysine (Lai and Gunsalus, 1992; Lai et al., 1991). Production of these compounds at the rates and levels needed to compensate for high salinity, along with active regulation of ion transport across the cell membrane, constitutes significant and ongoing energy expenditure. Indeed, the energy expenditure to maintain osmotic balance and regulate intracellular salt concentrations can be expected to increase monotonically with the salinity of the environment. Thus, salinity based limitations on methanogenic metabolism may act principally at a bioenergetic level (Oren, 1999, 2001). Among halotolerant or halophilic methanogens, those representing methylotrophic modes of metabolism (excluding acetate) are typically capable of growth or metabolism at higher salinities than H2- or acetate-utilizing organisms. Oren (2001) has proposed that the greater standard free energy yield available to methylotrophic methanogens may account for this observation, noting that higher energy yields may serve to balance the higher rates of energy expenditure required for life at higher salinities. While higher overall rates of metabolic energy production will certainly serve to balance higher cellular maintenance energies, some caution is warranted in this interpretation. Specifically, higher standard Gibbs free energy yields are, by themselves, only one determinant of the total metabolic energy yield, and the rate of substrate flux/consumption must also be considered. For example, a methanogen consuming acetate with a five fold lower Gibbs free energy yield than a methanogen consuming methylamine will nonetheless have a higher overall rate of metabolic energy production if it receives a ten fold higher flux of substrate. Importantly, however, methylotrophic substrates – some of which are among the breakdown products of osmoregulants – appear to be proportionately more abundant as methanogenic substrates in hypersaline settings. In combination with the greater free energy yields of methylotrophic methanogenesis, this may help to support the higher salinity tolerance of methylotrophic methanogens than their H2- or acetateutilizing counterparts. Regardless, energy balance appears to be a critical determinant of the methanogenic habitability of hypersaline environments.

4.5

Pressure

Elevated pressure is a characteristic of many methanogenic environments (e.g., sediments underlying the deep ocean basins), and methanogen activity is documented to pressures of 75 MPa, equivalent to >7000 m water depth (Miller et al., 1998). However, the difficulties associated with conducting physiological studies at very high pressures has limited our direct understanding of the tolerance and adaptations of methanogens to high pressures. Because biochemical and metabolic reactions occur in aqueous solution, and because the partial molar volume changes associated with aqueous reactions are typically quite low, high pressures have only minimal effects on the thermodynamics of methane production (See > Chapter 21, Vol. 1, Part 5). Similarly, significant biochemical changes (e.g., pressure denaturation of proteins) are expected to require pressures substantially greater than 100 MPa, the approximate pressure associated with the deepest ocean trenches (Silva and Weber, 1993). Instead, the most relevant effects of pressure are likely on the substrate transport (energy yielding)

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contribution to cellular energy balance, via enhanced solubility (and correspondingly greater transport potential) of gaseous substrates and products (e.g., H2).

5

Research Needs

Culture-based microbiology has given us a snapshot of the methanogen tolerance to environmental extremes, but it is not clear how to translate these observations into a realistic predictor of their population distribution and activity in the natural world, where these organisms may function in the context of syntrophic and competitive interactions, and may face energy limitation and multiple physicochemical extremes. Advances in a variety of areas will ultimately help to address this question. The control that comes with studies of pure cultures or isolates is of great value and should unquestionably be brought to bear in considering the ‘‘physiological ecology’’ of methanogenesis. Traditionally, however, culture work optimizes energy availability and other growth factors, while seeking to isolate individual physicochemical variables for study. To begin to probe the question of survival in complex environments, these studies must begin to incorporate constraints – for example, energy limitation or combinations of physicochemical extremes – that present an increasingly realistic picture of the environment. Key targets of these studies should be to document the response of microbial energy metabolism (e.g., maintenance energy or compensating mechanisms) (Valentine, 2007) to deviations from optimal conditions and to better understand the biochemical and regulatory mechanisms associated with adaptation to extremes. Such studies will also benefit from continued attempts to sample the full diversity of methanogens in environments presenting individual and combinations of extremes (including energy limitation), to ensure that the full range of environmental tolerances and mechanisms of adaptation are reflected in cultured organisms. Beyond the petri dish, advances in characterizing microbial ecology and physiology in situ will significantly enhance our predictive capability regarding the environmental distribution of methanogenic activity vis-a`-vis a variety of extremes. Key areas for advancement include accurate in situ rate measurements, especially in cases of low metabolic activity (e.g., in cold or low energy settings); methods for discerning and discriminating metabolic status (e.g., active growth vs. simple maintenance) and for obtaining accurate cell counts at low numbers; methods for linking geochemical function with genetic identity; and means for better resolving complex ecological interactions and associations (including the syntrophic partnerships and competitive interactions with which methanogens in many environments are so inextricably linked). Finally, theoretical work on microbial energy metabolism may aid in developing a quantitative framework in which to understand and predict the effects of multiple environmental forcing factors. Such work should combine with culture based and environmental studies to assess the biochemical and bioenergetic impacts and adaptations associated with environmental extremes. Identifying common denominators (e.g., effects of diverse physicochemical extremes on cellular maintenance energies) will lead to an improved ability to quantify and predict compound effects on methanogen metabolism. Additionally, the development of numerical models for biological processes occurring at spatial or temporal scales that are not currently accessible by experimental or observational means (e.g., substrate transport and reaction at single-cell scales) will impose quantitative constraints on questions that currently constitute a significant knowledge gap.

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Acknowledgments This work was supported in part by awards DE-FG02-96ER20214 and DE-FG02-08ER64689 from the Department of Energy and NSF EF-0333294 from the National Science Foundation.

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Methanococcoides burtonii. Genome Res 13: 1580–1588. Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61: 262–280. Schink B (1988) Principles and limits of anaerobic degradation. In Biology of Anaerobic Microorganisms. AJB Zehnder (ed.). New York: Wiley, pp. 771–846. Schink B, Stams AJM (2006) Syntrophism among prokaryotes. In The Prokaryotes: An Evolving Electronic Resource for the Microbiological Community, 3rd edn, Vol. 2. M Dworkin, S Falkow, E Rosenberg, KH Schleifer, and E Stackebrandt (eds.). New York: Springer, pp. 309–335. Sekiguchi Y, Kamagata Y, Nakamura K, Ohashi A, Harada H (2000) Syntrophothermus lipocalidus gen. nov., sp. nov., a novel thermophilic, syntrophic, fatty-acid-oxidizing anaerobe which utilizes isobutyrate. Int J Syst Evol Microbiol 50: 771–779. Shock EL, Holland ME (2007) Quantitative habitability. Astrobiology 7: 839–851. Silva JL, Weber G (1993) Pressure stability of proteins. Annu Rev Phys Chem 44: 89–113. Simankova MV, Kotsyurbenko OR, Lueders T, Nozhevnikova AN, Wagner B, Conrad R, Friedrich MW (2003) Isolation and characterization of new strains of methanogens from cold terrestrial habitats. Syst Appl Microbiol 26: 312–318. Simankova MV, Parshina SN, Tourova TP, Kolganova TV, Zehnder AB, Nozhevnikova AN (2001) Methanosarcina lacustris sp. nov., a new psychrotolerant methanogenic archaeon from anoxic lake sediments. Syst Appl Microbiol 24: 362–367. Stetter KO, Thomm M, Winter J, Wildgruber G, Huber H, Zillig W, Jane´covic D, Ko¨nig H, Palm P, Wunderl S (1981) Methanothermus fervidus, sp. nov., a novel extremely thermophilic methanogen isolated from an Icelandic hot spring. Zentralbl Bakteriol Parasitenkd Infektionskr Hyg Abt 1 Orig Reihe C2: 166–178. Svetlitshnyi V, Rainey F, Wiegel J (1996) Thermosyntropha lipolytica gen. nov., sp. nov., a lipolytic, anaerobic, alkalitolerant, thermophilic bacterium utilizing short- and long-chain fatty acids in syntrophic coculture with a methanogenic archaeum. Int J Syst Bacteriol 46: 1131–1137. Takai K, Nakamura K, Toki T, Tsunogai U, Miyazaki M, Miyazaki J, Hirayama H, Nakagawa S, Nunoura T, Horikoshi K (2008) Cell proliferation at 122
C and isotopically heavy methane under high-pressure cultivation. Proc Natl Acad Sci USA 105: 10949–10954.

Takai K, Nealson KN, Horikoshi K (2004) Methanotorris formicicus sp. nov., a novel extremely thermophilic methane-producing archaeon isolated from a black smoker in Central Indian Ridge. Int J Syst Bacteriol 54: 1095–1100. Thamdrup B (2000) Bacterial manganese and iron reduction in aquatic sediments. Adv Micro Ecol 16: 41–84. Tijhuis L, van Loosdrecht MCM, Heijnen JJ (1993) A thermodynamically based correlation for maintenance Gibbs energy requirements in aerobic and anaerobic chemotrophic growth. Biotechnol Bioeng 42: 509–519. Ueno Y, Yamada K, Yoshida N, Maruyama S, Isozaki Y (2006) Evidence from fluid inclusions for microbial methanogenesis in the early Archaean era. Nature 440: 516–519. Valentine DL (2007) Adaptations to energy stress dictate the ecology and evolution of the Archaea. Nature Rev Microbiol 5: 316–323. von Klein D, Arab H, Vo¨lker H, Thomm M (2002) Methanosarcina baltica , sp. nov., a novel methanogen isolated from the Gotland Deep of the Baltic Sea. Extremophiles 6: 103–110. Wilhelms A, Larter SR, Head I, Farrimond P, di-Primio R, Zwach C (2001) Biodegradation of oil in uplifted basins prevented by deep-burial sterilization. Nature 411: 1034–1037. Williams RT, Crawford RL (1984) Methane production in Minnesota peatlands. Appl Environ Microbiol 47: 1266–1271. Wolfe RS (1971) Microbial formation of methane. Adv Microb Phys 6: 107–146. Wolin MJ, Miller TL (1982) Interspecies hydrogen transfer: 15 years later. ASM News 48: 561–565. Yu IK, Kawamura F (1987) Halomethanococcus doii gen. nov., sp. nov.: an obligately halophilic methanogenic bacterium from solar salt ponds. J Gen Appl Microbiol 33: 303–310. Zehnder AJB, Stumm W (1988) Geochemistry and biogeochemistry of anaerobic habitats. In Biology of Anaerobic Microorganisms. AJB Zehnder (ed.). New York: Wiley, pp. 1–38. Zinder S (1993) Physiological ecology of methanogens. In Methanogenesis, JF Ferry (ed.). New York: Chapman and Hall, pp. 128–206. Zhilina TH (1983) A. new obligate halophilic methaneproducing bacterium. Mikrobiologiya 52: 375–382. Zhilina TN, Zavarzin GA (1987) Methanohalobium evestigatus, gen. nov., sp. nov., the extremely halophilic methanogenic archaebacterium. Dokl Akad Nauk SSSR 293: 464–468.

52 Methanogenesis in Arctic Permafrost Habitats D. Wagner1 . S. Liebner 2 1 Alfred Wegener Institute for Polar and Marine Research, Research Unit Potsdam, Potsdam, Germany [email protected] 2 Institute for Biogeochemistry and Pollutant Dynamics (IBP), Federal Institute of Technology (ETH), Universita¨tstrasse, Zu¨rich, Switzerland [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 656 2 The Permafrost Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 656 3 Methane Cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 657 4 Diversity and Ecology of Methanogenic Archaea in Permafrost Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 658 5 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 661

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_52, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: In polar regions, huge layers of frozen ground, termed permafrost, are formed. Permafrost covers more than 25% of the land surface and significant parts of the coastal sea shelves. Permafrost habitats are controlled by extreme climate and terrain conditions. Particularly, the seasonal freezing and thawing in the upper active layer of permafrost leads to distinct gradients in temperature and geochemistry. Methanogenic archaea in permafrost environments have to survive extremely cold temperatures, freeze-thaw cycles, desiccation and starvation under long-lasting background radiation over geological time scales. Although the biology of permafrost microorganisms remains relatively unexplored, recent findings show that methanogenic communities in this extreme environment are composed by members of the major phyla of the methanogenic archaea (Methanobrevibacter, Methanobacterium, Methanosaeta, Methanosarcina, Methanolobus/Methanohalophylus/Methanococcoides, Methanoculleus/Methanogenium), with a total biomass comparable to temperate soil ecosystems. Currently, methanogenic archaea were the object of particular attention in permafrost studies, because of their key role in the Arctic methane cycle and consequently of their significance for the global methane budget.

1

Introduction

The Arctic plays a key role in the Earth’s climate system for two reasons. On the one hand, global warming is predicted to be most pronounced at high latitudes, and observational evidence over the past 25 years suggests that this warming is already under way (RichterMenge et al., 2006). On the other hand, one third of the global carbon pool is stored in ecosystems of the northern latitudes (Post et al., 1982; Zimov et al., 2006). Thus there is considerable socio-economic interest in predicting how the carbon balance of the northern ecosystems will respond to ongoing climate warming. The degradation of permafrost and the associated intensified release of methane, a climaterelevant trace gas, represent potential environmental hazards (Anisomov et al., 1999). The microorganisms driving anaerobic carbon decomposition processes including methane production in Arctic permafrost environments have remained poorly investigated. Their population structure and reaction to environmental changes is largely unknown, which means that also an important part of the process knowledge on greenhouse gas fluxes in permafrost ecosystems is far from completely understood. This also hampers prediction of the effects of climate warming on Arctic methane fluxes. Further research on the stability of the methane cycling communities is therefore highly important for understanding the effects of a warming Arctic on the global climate (See also > Chapter 40, Vol. 3, Part 3). This chapter first gives an introduction into permafrost as a habitat for microorganisms. It then describes the current knowledge on the diversity and ecology of methane-producing archaea and gives an outlook for further research needs.

2

The Permafrost Environment

Permafrost, which covers about 25% of the Earth’s land surface (Zhang et al., 1999) and significant parts of the coastal sea shelfs (Romanovskii et al., 2005), is defined as ground, comprised of soil or rock and included ice and organic material that remain at or below 0 C for

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at least two consecutive years (van Everdingen, 2005). Arctic permafrost regions are characterized by low mean annual air temperatures (from –8 to –15 C), low mean annual precipitations (from 90 to 370 mm) and poor to missing vegetation (French, 1996; ROSHYDROMET, 2004). During the relatively short period of arctic summer, only the surface zone (a few dm-thick) of permafrost sediments thaws: this is called the active layer. Active layer depths range from a few cm in the high Arctic to more than 2 m in subarctic regions. Permafrost can be cemented by ice, which is typical for Arctic regions, or, in the case of insufficient interstitial water, may be dry as occurs in the Antarctic polar deserts or rocky areas. The boundary between the active layer and the perennially frozen ground is called the permafrost table, which acts as a physical and chemical barrier. Intensive physico-chemical processes under extreme conditions take place in the active layer and upper permafrost sediments (Ostroumov, 2004). In the deeper permafrost layers, conditions have been stable for long periods of time and microbial processes are limited (Wagner, 2008). Permafrost soils (cryosols) have been developed in the upper zone of the cryolithosphere (active layer and upper permafrost sediments) where the temperatures range from –50 C to + 30 C (Yershov, 1998). Therefore, permafrost soils are mainly formed by cryopedogenesis, which involves freezing and thawing, frost stirring, mounding, fissuring and solifluction. The repeating cycles of freezing and thawing leads to cryoturbation features (frost churning) that includes irregular, broken or involuted horizons and an enrichment in organic matter and inorganic compounds, especially on top of the permafrost table (Bockheim et al., 1999; Van Vliet-Lanoe¨, 1991). As a result of cryopedogenesis, many permafrost soils are influenced by a strong micro-relief (e.g., low-centered ice-wedge polygons), which causes small-scale variations in soil types and vegetation characteristics (Kutzbach et al., 2004), as well as in the microclimatic conditions (Boike et al., 2008). This affects the abundance, processes and diversity of the methanogenic community in this habitat.

3

Methane Cycle

The carbon pool estimates for permafrost soils vary between 190 Gt and, in more recent studies, approximately 900 Gt (Post et al., 1982; Anisimov and Reneva 2006; Zimov et al., 2006). These large variations can be attributed to different soil types (from mineral to peaty soils) and varying depths for the calculation (from the upper few centimeters to several m depth). The degradation of permafrost, therefore, could release large quantities of previously frozen organic matter. Permafrost degradation through environmental changes is considered to have a stronger impact on organic carbon decomposition rates than the direct effect of temperature rise alone (Eugster et al., 2000). This process is associated with the release of climate-relevant trace gases from intensified microbial carbon turnover that may further increase global warming and transform the Arctic tundra ecosystems from a carbon sink to a carbon source (Oechel et al., 1993). However, permafrost soils can function as both a source and a sink for carbon dioxide and methane. Under anaerobic conditions, caused by flooding of the permafrost soils and the effect of backwater above the permafrost table, the mineralization of organic matter can only be realized stepwise by specialized microorganisms of the so called anaerobic food chain (Schink and Stams, 2006). Important intermediates of the organic matter decomposition under anaerobic conditions are polysaccharides, low-molecular-weight organic acids,

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phenolic compounds and sugar monomers (Guggenberger et al., 1994; Kaiser et al., 2001). These compounds will be further converted for instance into hydrogen, carbon dioxide and acetate, which can be reduced to methane (methanogenesis) by methanogenic archaea. The fermentation of carbon by microorganisms runs much slower than the oxidative respiration. As a result of the prolonged anaerobic conditions and low in situ temperatures of permafrost soils organic matter accumulates (peat formation) in these environments (sea above). Wherever oxygen is present in permafrost habitats (upper toxic soil horizons, rhizosphere) methane can be oxidized to carbon dioxide by aerobic methane oxidizing bacteria (for details See > Chapter 40, Vol. 3, Part 3).

4

Diversity and Ecology of Methanogenic Archaea in Permafrost Environments

Responsible for the biogenic methane production (methanogenesis) is a small group of microorganisms called methanogenic archaea (Garcia, 1990). They can be found either in temperate habitats like paddy fields (Grosskopf et al., 1998), lakes (Jurgens et al., 2000; Keough et al., 2003), freshwater sediments (Chan et al., 2005), in the gastrointestinal tract of animals (Lin et al., 1997), or in extreme habitats such as hydrothermal vents (Jeanthon et al., 1999), hypersaline habitats (Mathrani and Boone, 1985) or permafrost soils and sediments (Kobabe et al., 2004; Rivkina et al., 1998). Although methanogens are widely spread in nature they show an extremely specialized metabolism. They are able to converse only a limited number of substrates (e.g., hydrogen, acetate, formate, methanol, methylated amines) to methane. In cold environments such as permafrost two main pathways of energy-metabolism dominate: (1) the reduction of CO2 to CH4 using H2 as a reductant and (2) the fermentation of acetate to CH4 and CO2. In the case of CO2-reduction organic carbon is not necessary for growth of methanogenic archaea (Conrad, 2005). At present, 26 genera with altogether 107 species of methanogenic archaea are described (http://www.ncbi.nlm.nih.gov/taxonomy). Genera with the most described species are Methanobacterium, Methanobrevibacter, and Methanosarcina (number of species between 9 and 14). Phylogenetically, they are classified as Archaea (Whitman et al., 2006), a group of microbes that are distinguished from Bacteria by some specific characteristics (e.g., cell wall composition, coenzymes). Methanogenic archaea are widespread in nature and highly abundant in extreme environments tolerating low/high temperatures (permafrost, hot springs), extreme salinity (saltern ponds) and low/high pH (solfataras, soda lakes). Although, they are regarded as strictly anaerobic organisms without the ability to form spores or other resting stages, they are found in millions of years old permafrost sediments (Shi et al., 1997). In addition to mesophilic species, thermophilic and hyperthermophilic methanogens have also been identified (Garcia et al., 2000; Stetter et al., 1990). Recently, more attention has been paid to the isolation of psychrophilic strains since a number of methanogenic habitats are located in cold climates (Gounot, 1999). Although the metabolism of methanogenic archaea has been studied in different environments (Eicher, 2001; Garcia et al., 2000; Lange and Ahring, 2001; Shuisong and Boone, 1998), only a few studies have focussed on the ecology of the methanogenic archaea exposed to the harsh environmental conditions of permafrost, e.g., subzero temperatures, low water activity and low nutrient availability (Ganzert et al., 2007; Høj et al., 2005; Vishnivetskaya et al., 2000).

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Although permafrost environments are characterized by extreme climate conditions, it was recently shown that the abundance and composition of the methanogenic population is similar to that of communities of comparable temperate soil ecosystems (Wagner et al., 2005). The highest cell counts of methanogenic archaea were detected in the active layer of permafrost, with numbers of up to 3  108 cells g 1 soil (Kobabe et al., 2004). Methanogenic archaea represented between 0.5 and 22.4% of the total cell counts. Phylogenetic analyses revealed a great diversity of methanogens in the active layer, with species belonging to the families Methanobacteriaceae, Methanomicrobiaceae, Methanosarcinaceae, and Methanosaetaceae (Høj et al., 2005; Ganzert et al., 2007; Metje and Frenzel, 2007). Other sequences detected were affiliated to the euryarchaeotal Rice Cluster II and V (Grosskopf et al., 1998; Hales et al., 1996; Ramakrishnan et al., 2001) as well as to the Group I.3b of the uncultured Crenarchaeota (nonmethanogenic archaea; Ochsenreiter et al., 2003). Environmental sequences from the Laptev Sea coast form four specific permafrost clusters (Ganzert et al., 2007). Permafrost Cluster I was recovered mainly from cold horizons (with temperatures of less than 4 C) of the active layer and was related to Methanosarcinacea. Permafrost Clusters II and III were related to Methanomicrobiales and Permafrost Cluster IV was related to Rice Cluster II. It was hypothesized that these clusters comprise methanogenic archaea with a specific physiological potential to survive under harsh environmental conditions. The phylogenetic affiliation of the sequences recovered in the study by Ganzert and colleagues, (2007) indicated that both hydrogenotrophic and acetoclastic methanogenesis exist in permafrost soils. Recent studies on perennially frozen permafrost deposits from the Lena Delta (Siberia) revealed significant amounts of methane which could be attributed to in situ activity of methanogenic archaea (Wagner et al., 2007). Another study on frozen ground on Ellesmere Island reported an archaeal community composed of 61% Euryarchaeota (i.e., methane producing archaea) and 39% Crenarchaeota, suggesting the presence of a diverse archaeal population also in the perennially frozen sediments (Steven et al., 2007). First studies on submarine permafrost sediments indicate a different methanogenic community in comparison to its terrestrial counterpart (Koch et al., 2009). Samples with high methane concentrations were dominated by sequences affiliated to the methylotrophic genera Methanosarcina and Methanococcoides as well as uncultured archaea. So far, only a few psychrophilic and psychrotrophic strains as well as several uncultivated methanogens were obtained from Arctic and Antarctic habitats (> Fig. 1): Methanococcoides burtonii was isolated from the anoxic hypolimnion of the Ace Lake, Antarctica (Franzmann et al., 1992) and Methanococcoides alaskense was obtained from marine sediments from Skan Bay, Alaska (Singh et al., 2005). Both organisms are cold-adapted with a minimum temperature for growth of 1.7 and –2.3, respectively. Cells grew with trimethylamine as a catabolic substrate and some strains could also grow with methanol. Methanogenium frigidum was also isolated from the Ace Lake, Antarctica (Franzmann et al., 1997). The cells exhibiting most rapid growth at 15 C and no growth at temperatures above 20 C. The organisms grew by CO2 reduction by using H2 as a reductant. Formate could replace H2, while acetate, methanol and trimethylamine were not catabolized. Methanosarcina spec. SMA-21, which is closely related to Methanosarcina mazei, was recently isolated from a Siberian permafrost soil in the Lena Delta. The organism grows well at 28 C and slowly at low temperatures (4 and 10 C) with H2/CO2 as substrate. The cells grow as cocci, with a diameter of 1–2 mm. Methanosarcina SMA-21 is characterized by an extreme tolerance to very low temperatures (–78.5 C), high salinity (up to 6 M), starvation, desiccation and oxygen exposure (Morozova and Wagner, 2007). In addition, this archaeon survived for 3 weeks under simulated thermo-physical

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. Figure 1 Phylogenetic relation (based on 16S rRNA gene sequences) of methanogenic archaea. Grey squares illustrate sequences with Arctic tundra origin (or groups containing sequences from Arctic tundra environments). Tree represent a maximum likelihood tree and were constructed using the ARB software package.

Martian conditions (Morozova et al., 2007). Furthermore, five new strains of methanogenic archaea were isolated from permanently or periodically cold terrestrial habitats in Russia and Switzerland (Simankova et al., 2003). Three of them were members of the methylotrophic genus Methanosarcina, one hydrogenotrophic strain is a new ecotype of the genus Methanocorpusculum and one obligately methylotrophic strain is closely related to Methanomethylovorans hollandica. All new isolates are not true psychrophiles according to their growth temperature characteristics. In spite of the ability of all isolates to grow at temperatures as low as 1–5 C, all of them have their growth optima in the range of moderate temperatures (25–35 C). Thus, they can be regarded as psychrotolerant organisms. Methane production was observed at low in situ temperatures with rates of up to 39 nmol CH4 h 1 g 1 soil in the active layer of permafrost (Høj et al., 2005; Metje and Frenzel, 2007; Wagner et al., 2003). The highest activities were thereby measured in the coldest zones of the profiles. Furthermore, it could be shown that methane production is rather limited by the quality of soil organic carbon than by the in situ temperature (Ganzert et al., 2007; Wagner

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et al., 2005). Another important factor affecting methanogenic communities in permafrost soils is the water regime. Along a natural soil moisture gradient, changes in archaeal community composition were observed, which suggest that the differences in these communities were responsible for the large-scale variations in methane emissions observed with changes in soil hydrology (Høj et al., 2006; Wagner et al., 2003). The permafrost environment forces the adaptation of the microbial communities to low temperature conditions and promotes the growth of species that so far remain undetected in temperate ecosystems. Therefore, Arctic permafrost environments can be seen as active microbial ecosystems rather than frozen habitats with microbial survivors. The evaluation of microbiological data and their correlation with climatic and geochemical results represents the basis for the understanding of the role of permafrost in the global system. Of particular relevance are feedback mechanisms related to nutrient cycles, biogeochemical processes and greenhouse gas emissions in the context of a warming Earth.

5

Research Needs

Although one fourth of the Earth land surface and distinct areas of the coastal sea shelfs are affected by permafrost, the physiology, function and diversity of microbial communities in these ecosystems is sparsely investigated so far. This may be partially caused by the relative inaccessibility of the investigation areas and the associated logistic problems. However, the main difficulty lies in the lack of methodologies specific for permafrost sampling and isolation of cold-adapted microorganisms from Arctic soils and sediments. This is shown by the discrepancy between the small numbers of psychrophilic microorganism isolated so far from permafrost environments in contrast to the observed significant metabolic rates under in situ conditions. Methodological developments should consider the following aspects: enrichment of microorganisms should be performed directly in the field or in batch or continuous laboratory culture; culture techniques should be developed for the enrichment of ‘‘syntrophically associated’’ microorganisms; sub-zero culturing methods are needed; and state-of-the-art culture-independent molecular techniques for diversity and functional analyses of microbial communities should be applied on permafrost. The lack of isolates from permafrost limits also possible biotechnological uses. Coldadapted microorganisms from permafrost exhibit properties very different from those of other thermal classes. Therefore, the vast genetic resources of microorganisms from permafrost environments remain nearly unexploited. It is likely that mainly extremophilic microbes could offer technologically and/or economically significant products such as enzymes, polysaccharides, osmoprotectors and liposomes (Cavicchioli et al., 2002). Therefore, one essential goal of microbial diversity exploration in cold regions will be to recover new isolates, some of which will prove useful for biotechnology processes or medicine.

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Methanogenesis in Arctic Permafrost Habitats Metje M, Frenzel P (2007) Methanogenesis and methanogenic pathways in a peat from subarctic permafrost. Environ Microbiol 9: 954–964. Ochsenreiter T, Selezi D, Quaiser A, BonchOsmolovskaya L, Schleper C (2003) Diversity and abundance of Crenarchaeota in terrestrial habitats studied by 16S RNA surveys and real time PCR. Environ Microbiol 5: 787–797. Oechel WC, Hastings SJ, Jenkins M, Riechers G, Grulke NE, Vourlitis GL (1993) Recent change of arctic tundra ecosystems from a net carbon sink to a source. Nature 361: 520–526. Ostroumov V (2004) Physico-chemical processes in cryogenic soils. In Cryosols. JM Kimble (ed.). Berlin: Springer, pp. 347–364. Post WM, Emanuel WR, Zinke PJ, Stangenberger AG (1982) Soil carbon pools and world life zones. Nature 298: 156–159. Ramakrishnan B, Lueders T, Dunfield PF, Conrad R, Friedrich MW (2001) Archaeal community structures in rice soils from different geographical regions before and after initiation of methane production. FEMS Microbiol Ecol 37: 175–186. Richter-Menge J, Overland J, Proshutinsky A, et al. (2006) State of the Arctic Report. NOAA OAR Special Report, NOAA/OAR/PMEL. Seattle, WA, p. 36. Rivkina EM, Gilichinsky D, Wagener S, Tiedje J, McGrath J (1998) Biochemical activity of anaerobic microorganisms from buried permafrost sediments, Geomicrobiol J 15: 187–193. Romanovskii NN, Hubberten H-W, Gavrilov AV, Eliseeva AA, Tipenko GS (2005) Offshore permafrost and gas hydrate stability zone on the shelf of East Siberian Seas. Geo-Marine Lett 25: 167–182. ROSHYDROMET (2004) Russian Federal Service for Hydrometeorology and Environmental Monitoring, http://www.worldweather.org/107/c01040.htm. Schink B, Stams AJM (2006) Syntrophism among Prokaryotes. In Prokaryotes, vol 2. M Dworkin, S Falkow, E Rosenberg, K-H Schleifer, E Stackebrandt (eds.). New York: Springer, pp. 309–335. Simankova MV, Kotsyurbenko OR, Lueders T, Nozhevnikova AN, Wagner B, Conrad R, Friedrich MW (2003) Isolation and characterization of new strains of methanogens from cold terrestrial habitats. Syst Appl Microbiol 26: 312–318. Singh N, Kendall MM, Liu Y, Boone DR (2005) Isolation and characterization of methylotrophic methanogens from anoxic marine sediments in Skan Bay, Alaska: description of Methanococcoides alaskense sp. nov., and emended description of Methanosarcina baltica. Int J Syst Evol Microbiol 55: 2531–2538. Shi T, Reevers R, Gilichinsky D, Friedmann EI (1997) Characterization of viable bacteria from Siberian permafrost by 16S rDNA sequencing. Microbial Ecol 33: 167–179.

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Shuisong N, Boone D (1998) Extremophilic methanogenic archaea and their adaptation mechanisms. In Extremophiles: Microbial Life in Extreme Environments, K Horikoshi and WD Grant (eds.). New York: Wiley, 211–232. Stetter KO, Fiala G, Huber G, Huber R, Segerer A (1990) Hyperthermophilic microorganisms. FEMS Microbiol Rev 75: 117–124. Steven B, Briggs G, McKay CP, Pollard WH, Greer CW, Whyte LG (2007) Characterization of the microbial diversity in a permafrost sample from the Canadian high Arctic using culture-dependent and cultureindependent methods. FEMS Microbiol Ecol 59: 513–523. Van Everdingen R (2005) Multi-language glossary of permafrost and related ground-ice terms. Boulder, CO: National Snow and Ice Data Center/World Data Center for Glaciology. Van Vliet-Lanoe¨ B. (1991) Differential frost heave, load casting and convection: converging mechanisms; a discussion of the origin of cryoturbations. Permafrost Periglac Process 2: 123–139. Vishnivetskaya T, Kathariou S, McGrath J, Gilichinsky D, Tiedje J (2000) Low-temperature recovery strategies for the isolation of bacteria from ancient permafrost sediments. Extremophiles 4: 165–173. Wagner D, Kobabe S, Pfeiffer EM, Hubberten (2003) Microbial controls on methane fluxes from a polygonal tundra of the Lena Delta, Siberia. Permafrost Periglac Process 14: 173–185. Wagner D, Lipski A, Embacher A, Gattinger A (2005) Methane fluxes in extreme permafrost habitats of the Lena Delta: effects of microbial community structure and organic matter quality. Environ Microbiol 7: 1582–1592. Wagner D, Gattinger A, Embacher A, Pfeiffer EM, Schloter M, Lipski A (2007) Methanogenic activity and biomass in Holocene permafrost deposits of the Lena Delta, Siberian Arctic and its implication for the global methane budget. Global Change Biol 13: 1089–1099. Wagner D (2008) Microbial communities and processes in Arctic permafrost environments. In Microbiology of extreme soils, Soil Biology, vol. 13., P Dion and CS Nautiyal (eds.). Berlin: Springer, 133–154. Whitman WB, Bowen TL, Boone DR (2006) The methanogenic bacteria. Prokaryotes 3: 165–207. Yershov ED (1998) General geochryology. Cambridge: Cambridge University Press. Zhang T, Barry RG, Knowles K, Heginbotton JA, Brown J (1999) Statistics and characteristics of permafrost and ground-ice distribution in the northern hemisphere. Polar Geography 23: 132–154. Zimov SA, Schuur EAG, Chapin III FS (2006) Permafrost and the global carbon budget. Science 312: 1612–1613.

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53 Methanogens and Methanogenesis in Hypersaline Environments T. J. McGenity Department of Biological Sciences, University of Essex, Colchester, UK [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 666 2 Sedimentary Rocks . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 668 3 Deep-Sea Hypersaline Anoxic Brine Lakes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 670 4 Guerrero Negro and Other Hypersaline Microbial Mats . . . . . . . . . . . . . . . . . . . . . . . . . . . . 671 5 Buried Salt Deposits . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 673 6 Other Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 673 7 Cycling of Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 674 8 Cultivated Halophilic Methanogens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 675 9 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 676

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Abstract: Methanogenesis in hypersaline environments is determined by redox potential and permanency of anaerobic conditions, and by the concentration of other terminal electron acceptors, particularly sulfate, because sulfate-reducing bacteria have a greater affinity than methanogens for competitive substrates like hydrogen and acetate. Hypersalinity, however, is not an obstacle to methanogenesis; in many cases it provides higher concentrations of noncompetitive substrates like methylamines, which derive from compatible solutes such as glycine-betaine that is synthesized by many microbes inhabiting hypersaline environments. Also, depletion of sulfate, as may occur in deeper sediments, allows increased methanogenesis. On the other hand, increasing salinity requires methanogens to synthesize or take up more compatible solutes at a significant energetic cost. Aceticlastic and hydrogenotrophic methanogens, with their lower energetic yields, are therefore more susceptible than methylotrophic methanogenesis, which further explains the predominance of methylotrophic methanogens like Methanohalophilus spp. in hypersaline environments. There are often deviations from the picture outlined above, which are sometimes difficult to explain. Identifying the role of uncultivated Euryarchaeota in hypersaline environments, elucidating the effects of different ions (which have differential stress effects and potential as electron acceptors) and understanding the effects of salinity on all microbes involved in methane cycling, will help us to understand and predict methane production in hypersaline environments.

1

Introduction

Hypersaline environments are simply defined as those with a greater concentration of salts than seawater. Such environments are many and varied, in terms of their overall salinity and predominant ions. Coastal environments, both man-made and natural are subject to desiccation, resulting in a wide variety of habitats from small, ephemeral salt pans within temperate salt marshes to large, permanently hypersaline sabkhas in sub-tropical regions (Hovorka, 1987). Similarly, inland salt lakes can be as large as the Great Salt Lake or a tiny spring. Salt deposits, often several hundred meters in thickness, lie beneath about a quarter of the Earth’s landmass, and contain brines from a cubic micrometer in volume to many cubic meters. Hypersaline environments are widespread and were more prevalent in former geological times, for example much of northern Europe was covered by the salt-saturated Zechstein Sea during the Permo-Triassic period (Zharkov, 1981), and the Mediterranean Sea was desiccated more recently, with halite precipitation starting between 5.6 and 5.55 million years ago (Hsu¨ et al., 1973). Deep-sea, anoxic, hypersaline brines, derived from dissolution of such ancient evaporites, form large lakes on the floor of the Gulf of Mexico, Mediterranean and Red Sea. Oil reservoirs are frequently associated with hypersaline environments, and many industrial waste streams are both anaerobic and hypersaline. The inhabitants of hypersaline environments are generally termed halophiles, and the use and misuse of this term, together with all its qualifiers, as well as examples of the most ecologically important extreme halophiles, have been discussed by Oren (2008). The ability of microbes to tolerate hypersaline environments with different chemical compositions varies widely: Don Juan Pond, a CaCl2-saturated brine, appears to support no life (see Oren, 2002), whereas the African soda lakes are amongst the most productive environments in the world (Grant and Tindall, 1986). Salinity was found to be the main factor influencing microbial community composition in a recent synthesis of 111 studies (Lozupone and Knight, 2007),

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and contrary to popular perception, microbial diversity can be extremely high in environments where the salinity is about two to three times higher than seawater and where redox and light gradients exist. For example, 42 of the main bacterial phyla and 15 novel candidate phyla were reported in a microbial mat in Guerrero Negro with a salinity of 8% (Ley et al., 2006), representing an unprecedented level of microbial diversity (Lozupone and Knight, 2007). Also, where salinity gradients occur, microbial biodiversity, abundance and activity can also be greatly elevated owing to a cocktail of electron acceptors, donors, nutrients and carbon sources in the ionic and redox gradient (Daffonchio et al., 2006). In most environments methanogens are in competition with sulfate-reducing bacteria for the products of fermentation, particularly hydrogen and acetate, and it is well documented that where sulfate concentrations are sufficiently high, e.g., saline and hypersaline environments, sulfate reduction will be the dominant terminal-electron-accepting process owing to the higher affinity for these competitive growth substrates (e.g., Lovley et al., 1982). Nevertheless, methanogenesis remains an important process in marine and hypersaline environments, for example in sulfate-depleted zones in deeper sediments (Wilms et al., 2007), in areas with elevated hydrogen production (Hoehler et al., 2001; Buckley et al., 2008), and where carbon sources are available that cannot be used by sulfate reducers (Oremland et al., 1982; Winfrey and Ward, 1983). Such non-competitive substrates include methanol, dimethylsulfide and methylated amines, of which the latter derive from salinity-induced compatible solutes. A scheme for the production of methylamines is indicated in > Fig. 1. Dimethylsulfide derives primarily from hydrolysis of its precursor molecule dimethylsulfoniopropionate, which, like glycine betaine, is a compatible solute, and so is common in saline and hypersaline environments (Kiene et al., 1986; Kiene and Visscher, 1987). Therefore methylotrophic

. Figure 1 Central importance of trimethylamine for methanogenesis in hypersaline environments (adapted from Welsh (2000), with permission from Wiley-Blackwell). Only the main pathways are shown, and many other scenarios have been illustrated and described by Oren (1990).

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methanogens are nearly always the dominant methanogens in hypersaline conditions, including salt-saturated environments (5.2 M NaCl), although there are exceptions (Oremland and King, 1989). The relative importance of methylotrophic methanogenesis is well illustrated by an analysis of the upper salinity (in parentheses) at which pure methanogenic cultures have been shown to grow with various substrates (Oren, 1999): methylamines (27%), hydrogen and carbon dioxide (12%) and acetate (4%). These salinities should not be considered as the upper limit of activity in situ, but indicative of the relative importance of these substrates at different salinities. Competition with sulfate reducers, and the consequent reduced pool of halophilic hydrogenotrophic and aceticlastic methanogens on which natural selection can act, partly explain the observed differences in salinity tolerance. However, this trend is also governed by the relative energy gain from different methanogenic reactions per mole of substrate (methylotrophic >> hydrogenotrophic  aceticlastic), especially since halophiles must expend a lot of energy to maintain an osmotically balanced and functional cytoplasm via the biosynthesis and/or uptake of organic compatible solutes, and/or uptake of potassium ions (Oren, 1999). This review will focus more on those long-term, large-scale hypersaline anoxic environments, which have been studied in much greater depth, and their locations are shown in > Fig. 2. The emphasis will be on contemporary work which builds on studies outlined in some excellent reviews (e.g., Oremland and King, 1989; Ollivier et al., 1994; Oren, 1999; Marvin diPasquale et al., 1999; Oren, 2002). In order to understand the role of methanogenesis in carbon cycling in hypersaline environments evidence will be drawn from methane fluxes from field measurements, methane production rates (often from slurry experiments), cultivation, and investigation of uncultivated methanogenic communities. Methanogens are frequently studied without cultivation, owing to a generally good correspondence between phylogeny and phenotype that is less typical in other groups. Also, the mcrA gene, coding for methyl coenzyme-M reductase subunit-A, has proven valuable for investigating methanogens, and there is good correspondence between the phylogenies obtained with this functional gene and phylogenetic markers like 16S rRNA. This has resulted in widespread application of these markers to investigate methanogens in hypersaline (and many other) environments. Of course this is not absolute, and there is uncertainty over the phenotype of uncultivated organisms giving rise to 16S rRNA sequences that cluster within the Euryarchaeota but outside of known methanogens.

2

Sedimentary Rocks

Waldron et al. (2007) exploited a natural salinity gradient from 8 mM to 3.5 M Cl in the subsurface Antrim Shale, rich in methane derived from biodegraded hydrocarbons, to understand the salinity constraints on different types of methanogenesis. Methanogenesis was an important process in the shales, owing to a lack of competition with other terminal-electronaccepting processes, and it was evident that methanogens were capable of a high level of activity at their in-situ salinities, with the exception of the 3.5 M brine, in which there was no methane production. It is difficult to ascertain the percentage salinity of the two most saline brines (2.3 and 3.5 M Cl ) because the Na+ concentrations are 1.1 and 1.4 M respectively, implying the presence of other cations (perhaps K+, Mg2+, Ca2+) that were not measured. Based on most-probable-number enrichments, there is a clear change in

. Figure 2 Locations of some of the main hypersaline environments where methanogensis has been studied.

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Methanogens and Methanogenesis in Hypersaline Environments

methanogenic processes, with a mixture of hydrogenotrophic, aceticlastic and methylotrophic methanogenesis (primarily hydrogenotrophic) at low salinities and a predominance of methylotrophic methanogenesis at higher salinities (Waldron et al., 2007). This is supported by 16S rRNA clone libraries, in which all clone sequences resembled those from Methanocorpusculum spp. at 12.8 mM Cl , while most resembled the methylotrophic Methanohalophilus spp. at 2.3 M Cl . The second most abundant group from the high-salinity well was closely related to Methanoplanus petrolearius, previously shown to tolerate 0.86 M Cl , and able to use a range of compounds including hydrogen and carbon dioxide (Ollivier et al., 1997), hinting that hydrogenotrophic methanogenesis may be possible at this very high salinity. Indeed Ollivier et al. (1998) isolated Methanocalculus haloterans from a hypersaline oil reservoir, a hydrogenotrophic methanogen capable of growth up to 12% NaCl (= 2 M Cl ). Perhaps, the long-term stability of the shale (>7000 years) and the absence of competing processes have enabled hydrogenotrophic methanogens to adapt to a higher salinity (Waldron et al., 2007).

3

Deep-Sea Hypersaline Anoxic Brine Lakes

There are numerous locations in the deep sea and the deep subsurface where dissolution of rock salt has resulted in hypersaline brine seeps and lakes, which are often associated with methane seeps. In the case of the deep-sea hypersaline brine lakes, the density gradient between the hypersaline brine and overlying seawater, coupled with weak currents at depth, restrict mixing, which results in the hypersaline brines becoming anoxic. Such pools and lakes have been discovered on the floor of the Gulf of Mexico, Red Sea and Mediterranean Sea. These hypersaline brines have commonly been shown to be a source of biogenic methane, often mixed with geogenic methane (Charlou et al., 2003; Joye et al., 2005). A small brine pool, called NR-1, in the Gulf of Mexico is even surrounded by dense beds of Bathymodiolus mussels which house chemoautotrophic bacteria presumably fed by sulfide or methane from the anoxic brine lake (MacDonald et al., 1990). In sediments fed by a sulfate-depleted hypersaline brine seep, Paull et al. (1985) found that the carbon in mussel tissue was isotopically light and hence presumably derived (via endosymbiotic methanotrophs) from brine-derived methane. Therefore, there is good evidence that food-webs are stimulated by reduced compounds derived from microbial activity in hypersaline brines analogous to hydrothermal vent communities (Martens et al., 1991). In hypersaline brines on the floor of the Red Sea, such as the Kebrit and Shaban Basins, there are hints of the presence of methanogens from archaeal biomarkers (Michaelis et al., 1990) and euryarchaeal 16S rRNA gene amplicons (Eder et al., 2002), both of which could derive from non-methanogenic Archaea. More direct evidence has come from four of the eastern Mediterranean hypersaline basins where van der Wielen et al. (2005) detected methane production rates (mM CH4 d 1) of 85.8 (Urania), 16.9 (l’Atalante), 4.2 (Bannock) and 2.6 (Discovery). The most abundant uncultivated archaeal clones (termed MSBL-1) in most of these basins (van der Wielen et al., 2005) and in a hydrothermal mud vent beneath Urania hypersaline brine (Yakimov et al., 2007a), branched most closely to methanogens. The 16S rRNA signature from a very similar organism was also found in the anoxic hypolimnion of shallow hypersaline Solar Lake (Cytryn et al., 2000) and sediments of hypersaline Lake Chaka (Jiang et al., 2006). MSBL-1 Archaea probably represent a novel order, but despite their phylogenetic affiliation we currently have no firm evidence that they are methanogenic. Yakimov et al. (2007b) found a change in the main archaeal community across the 2-meter halocline from oxic Mediterranean seawater (depth 3498.5 m) to almost

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NaCl-saturated, anoxic l’Atalante brine; with a group related to ANME-1 (putative anaerobic methane-oxidizing group) co-existing with the aforementioned MSBL-1 group near the top of the interface, i.e., where salinity is not extremely high but where oxygen is highly depleted. MSBL-1 was detected also in the brine, but Methanohalophilus-related 16S rRNA sequences were most abundant in the deeper parts of the halocline and in the hypersaline brine. Messenger RNA coding for methyl coenzyme-M reductase from Methanohalophilus spp. was detected in the l’Atalante hypersaline brine (Hallsworth et al., 2007), and we isolated strains from l’Atalante sediment with 99% 16S rRNA sequence similarity to Methanohalophilus mahii using sulfate-free medium and non-competitive growth substrates (Sass, Timmis, McGenity, unpublished). This suggests that methanogenesis is occurring throughout the hypersaline brine of l’Atalante basin, and is primarily mediated by Methanohalophilus spp. using methylamines. In Bannock basin, which is chemically similar to l’Atalante but located on the opposite side of the Mediterranean trench, extremely low methane production rates were observed throughout the halocline from deep Mediterranean seawater to the hypersaline brine, but this increased to 3.5 mM CH4 d 1 in the near-salt-saturated brine (Daffonchio et al., 2006). 16S rRNA sequences from Methanohalophilus spp. and relatives were not found in Bannock interface or brine, whereas MSBL-1 was detected in both and ANME-1 in the interface (van der Wielen et al., 2005; Daffonchio et al., 2006), raising interesting questions about the factors influencing the distribution of methanogens. Three of the eastern Mediterranean brine lakes are dominated by NaCl, but the Discovery brine is unusual in that it derives from the dissolution of bischofite and so is an almost pure, 5 molar MgCl2 brine, with a water activity (aw) of 0.37, the lowest recorded in a marine environment and far below the current known limit of life (0.605) (see Hallsworth et al., 2007). Moreover, MgCl2 destabilizes biological macromolecules at high concentrations, and above a concentration of 2.3 M (in the absence of compensating solutes) appears to be inhibitory to life (Hallsworth et al., 2007). In support of this notion, is the detection of mRNA, a highly labile indicator of active microbes, from sulfate reducers and methanogens, only in the upper half of the halocline from seawater to Discovery brine ( Fig. 1) which provide an energyrich compound that can be used by many halophilic and halotolerant methanogens, but not by sulfate-reducing bacteria. Additionally, competition may be diminished by sulfate being

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used up, e.g., at depth or by precipitation of sulfate minerals. Nevertheless, growth at very high salinities is energetically demanding, and so often serves to inhibit methanogenesis.

8

Cultivated Halophilic Methanogens

Cultivated, taxonomically described halophilic methanogens are shown in > Table 1, and in addition there are several interesting halophilic strains whose names have not been validly published, especially in the genera Methanohalophilus and Methanohalobium (see Zhilina, 2001). Strain OCM 283, from an oil-reservoir brine, has a specific requirement for calcium and magnesium ions, reflecting the ionic composition of the oil-field brine from which it was isolated (Obraztsova et al., 1988). It was originally named ‘‘Methanococcoides euhalobius’’ (Obraztsova et al., 1988), but based on its wide NaCl tolerance (1–14%, optimum 6%), growth with methanol and methylamines and 16S rRNA sequence comparison, it was proposed to transfer it to the genus Methanohalophilus (Davidova

. Table 1 Cultivated, taxonomically described halophilic methanogens Order Speciesa

Strain

Habitatb

NaCl NaCl rangec optc

Carbon sourcesd

Original publication

Methanosarcinales Methanohalophilus portucalensis

OCM 59

Saltern sediments, Portugal

2–25

3–12

Methanol, MA

Boone et al. (1993)

Methanohalophilus (Methanococcus) halophilus

OCM 160 = DSMZ Shark Bay, 3094 Australia, microbial mat

2–15

7–9

Methanol, MA

Zhilina (1983)

Methanohalophilus mahii

OCM 68 = ATCC 35705

Great Salt Lake, sediment

3–20

6–15

Methanol, MA

Paterek and Smith (1988)

Methanosalsum (Methanohalophilus) zhilinae

OCM 62 = DSMZ 4017

Bosa Lake (Wadi Natrun)

1–12

4

Methanol, MA, DMS

Mathrani et al. (1988)

Methanohalobium evestigatum

OCM 161 = DSMZ Sivash Lake, 3721 microbial mat

15–30

25

Methanol, MA

Zhilina and Zavarzin (1987)

OCM 470

0–12

5

H2 & CO2, Formate

Ollivier et al. (1998)

Methanomicrobiales Methanocalculus halotolerans a

Oil field, Alsace

Genus names given in brackets indicate basonyms Habitat from which the type strain was isolated c NaCl range in which the species can grow, and opt = optimal salt concentration for growth; concentration in % w/v d MA methylamines; DMS dimethylsulfide b

675

676

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et al., 1997), however this name has never been validated. A strain whose name was validly published as Halomethanococcus doii (Yu and Kawamura, 1987), but which probably belongs to the genus Methanohalophilus, has been lost (Boone, 2001). Nakatsugawa (1991) described strain NY-218 which grew optimally at 14–18% NaCl and grew well on methylamines and methanol, and moderately on acetate. Methanocalculus halotolerans is the most halotolerant strain known that uses hydrogen and carbon dioxide, as well as formate, as growth substrates (Ollivier et al., 1998). However, Pe´rez-Fillol et al. (1985) reported an extremely halophilic, methanogen that was both hydrogenotrophic and methylotrophic. The genus Methanolobus has several slightly halophilic strains, such as Methanolobus oregonensis DSMZ 5435, which grows up to 9% NaCl, which, in addition to using methanol and methylamines, can use dimethylsulfide and methanethiol (Liu et al., 1990). The combined properties of halophily, alkaliphily and the ability to convert methanethiol to methane by Methanolobus oregonensis and related species are actively being tested for desulfurization of petroleum and gas (van Leerdam et al., 2008). Methanogens cope with elevated salinity by the accumulation of organic compatible solutes, often together with potassium ions which serve to counter the charge of anionic solutes. Additionally, enhanced expression of the gene coding for ClpB, a chaperone of the AAA+ superfamily, was identified in response to hyper- and hypo-osmotic stress (Shih and Lai, 2007). However, addition of glycine betaine reduced expression of clpB upon hyperosmotic shock, indicating that it has an important role in protein protection at high salinity. The common compatible solute, glycine betaine can be synthesized and accumulated by many methanogens, but they also use a variety of uncommon solutes, for example Methanohalophilus spp. use Ne-acetyl-b-lysine, b-glutamine, La-glutamate and a-glucosylglycerate (Lai et al., 1991; Roberts, 2005). Thus, on the one hand, not only does glycine betaine serve as a source of carbon and energy after fermentation to methylamines by other organisms, it is also an important osmoprotectant. It would be interesting to learn the extent to which halophilic methanogens ‘monitor’ the concentration of dissolved glycine betaine in the environment, and adjust its uptake and metabolism accordingly, on the one hand taking advantage of a pre-made compatible solute, while on the other retaining sufficient for future growth.

9

Research Needs

Few halophilic methanogens have been isolated recently, and publication dates in > Table 1 indicate that none has been taxonomically described in the last decade. Also, the previous discussion highlights the presence of numerous Euryarchaeota that may be halophilic methanogens, but which remain to be cultivated. Bringing such organisms into culture should be a research priority, as was done recently for peat-bog methanogens, now represented by ‘Methanoregula boonei’ (Bra¨uer et al., 2006), and with a fully sequenced genome. Molecular biological tools are under-developed for methanogens, which, coupled with the paucity of genome sequences of halophilic methanogens, means that we are only just beginning to understand the generic and methanogen-specific features of adaptation to high salinity. Certainly, the potential interaction between compatible-solute and chaperone networks deserves further investigation (Shih and Lai, 2007). Cross-referencing clone libraries of mcrA and 16S rRNA sequences is valuable to detect trends such as determining which uncultivated methanogens are common to hypersaline environments. However, doing this

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53

manually is inefficient and, especially with the plethora of pyrosequencing data emerging, it is important to develop and apply software to study methanogen biogeography. For methanogens that evade cultivation, metagenomics, single-celled genomics, and stable-isotope probing, following 13C (e.g., from methylamines and other growth substrates) into lipids and nucleic acids, can provide a link between function and phylogeny. There are numerous other areas that need further research if we are to truly understand carbon cycling past, present and future: it is important to understand how methanogens cope with desiccation (from rice paddies to salt marshes) because not only must methanogens tolerate low water activity, but also increased exposure to oxygen. What are the constraints of growth at high salinity – energetic (e.g., cost of making compatible solutes), competition, nutrient uptake etc., and how do these affect methane producers and consumers over space and time? Research into these areas may help us to learn whether and how aerobic Halobacteriaceae evolved from strictly anaerobic methanogens, and direct the search for putative missing links between the two archaeal groups.

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54 Thermophilic Methanoarchaea Inhabiting Hot Ecosystems B. Ollivier* . J.-L. Cayol Laboratoire de Microbiologie IRD, UMR 180, Universite´s de Provence et de la Me´diterrane´e, ESIL, Marseille, France *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 682

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Importance of Thermophilic Methanoarchaea in the Breakdown of Organic Matter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 685

3 Methanoarchaea from Extreme Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 687 3.1 Oilfield Reservoirs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 687 3.2 Deep-Sea Hydrothermal Vents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 688 4

Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 689

5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 690

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_54, # Springer-Verlag Berlin Heidelberg, 2010

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Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

Abstract: Thermophilic methanoarchaea are grouped into five orders. They range from moderately thermophiles to hyperthermophiles, Methanoplanus kandleri being the most thermophilic methanogen known so far (maximum temperature for growth is 110 C). They inhabit a wide range of hot terrestrial and subterrestrial ecosystems, including thermal springs, oil reservoirs, and deep sea hydrothermal vents, but also digestors operating at high temperatures. Their contribution to complete organic matter oxidation through hydrogen and acetate metabolism has been clearly established in digestors, whereas hydrogen, produced from biotic and abiotic reactions, appears as the primary energy source within the other hot ecosystems. Despite a limited metabolic diversity, thermophilic methanoarchaea are of ecological significance, in particular, in the geomicrobiology of the deep hot biosphere.

1

Introduction

Greater awareness of the role of methane in enhancing the greenhouse effect from CO2 accumulation has prompted an increasing number of studies on the global distribution of methane in the earth’s atmosphere. In recent years, more and more attention has been paid to the taxonomy, ecology, metabolism, energy use, biochemistry, and molecular biology of the methane-producing methanoarchaea. These strict anaerobes of the domain Archaea, phylum Euryarchaeota, share a complex biochemistry for methane synthesis as part of their energy metabolism (Garcia et al., 2000; Liu and Whitman, 2008). They are widely distributed in nature where they generally occupy the terminal niche in the transfer of electrons generated by the anaerobic degradation of organic matter. An exception is geothermally heated sediments where H2, formed by high-temperature abiotic reactions, is the primary source of energy for most of these microorganisms. The distribution of methanoarchaea in natural environments depends entirely on their adaptation to various physical and chemical variables, including temperature, pH, and salinity. Methanoarchaea can grow at up to 110 C, and hence, moderate thermophiles and hyperthermophiles have been frequently isolated from hot habitats and further characterized (> Table 1). The thermophilic species are distributed among 15 genera (> Fig. 1); five hyperthermophilic species growing at temperatures higher than 80 C belong to the genus Methanocaldococcus, two to the genus Methanothermus and Methanotorris, and one to the genus Methanopyrus; the moderate thermophiles are essentially members of the orders Methanosarcinales (Methanosarcina, Methanohalobium, and Methanomethylovorans) and Methanomicrobiales (Methanolinea) and will not grow at temperatures above 60 C. The remaining methanoarchaea are considered to be strictly thermophilic microorganisms, with optimum temperatures for growth in the range 55–70 C (> Table 1; > Fig. 1). Thermophilic methanoarchaea inhabit thermal environments such as hot springs, submarine hydrothermal vents, oil reservoirs and composts. They can also thrive in digesters operating at high temperature. All these environments are known to be sites of active methanogenesis. We note that thermophilic methanoarchaea can also be recovered not only from hot habitats but also from freshwater sediments, rivers, and ponds, where their contribution to organic matter oxidation is most probably low (Wagner and Wiegel, 2008). Like mesophilic methanoarchaea, thermophilic methanoarchaea use a narrow range of substrates (Liu and Whitman, 2008). While most of them utilize exclusively hydrogen (and in some cases formate) [Methanothermobacter, Methanothermus, and Methanocaldococcus species in particular], some are specialized in converting methanol and methylated amines

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

54

. Table 1 Comparison of morphological and physiological characteristics of thermophilic methanoarchaea Temperature Species

Range Morphology ( C)

Optimum ( C)

Methanogenesis substrates Typical habitats

Methanobacteriales Methanobacteriaceae H2 only

Mud from a cattle pasture

H2, formate

Anaerobic digestor sludge

65

H2 only

Sewage sludge

38–70

55

H2, formate

Anaerobic digestor sludge

Rod

45–65

57

H2 only

Thermophilic digestor

Methanothermobacter thermautotrophicus

Irregular curved rods

40–75

65–70

H2 only

Sewage sludge

Methanothermobacter wolfeii

Slender curved rods

37–74

55–65

H2 only

Sewage sludge and hot springs

Methanothermus fervidus

Curved rods

60–97

83

H2 only

Volcanic hot spring

Methanothermus sociabilis

Slightly curved rods

55–97

88

H2 only

Continental solfatara fields

Methanobacterium thermaggregans

Regular rods

40–75

65

Methanothermobacter defluvii

Rod

38–70

60–65

Methanothermobacter marburgensis

Rod

45–70

Methanothermobacter thermoflexus

Rod

Methanothermobacter thermophilus

Methanothermaceae

Methanococcales Methanococcaceae Methanothermococcus okinawensis

Irregular cocci

40–75

60–65

H2, formate

Deep sea hydrothermal vents

Methanothermococcus thermolithotrophicus

Regular cocci

30–70

65

H2, formate

Geothermally heated sea sediments

Methanocaldococcus jannaschii

Irregular cocci

50–86

85

H2 only

Submarine hydrothermal vent

Methanocaldococcus indicus

Irregular cocci

51–86

85

H2 only

Deep-sea hydrothermal chimney

Methanocaldococcus fervens

Irregular cocci

48–92

85

H2 only

Deep-sea hydrothermal chimney

Methanocaldococcaceae

683

684

54

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

. Table 1 (Continued) Temperature Optimum ( C)

Species

Range Morphology ( C)

Methanogenesis substrates Typical habitats

Methanocaldococcus infernus

Irregular cocci

55–91

85

H2 only

Deep-sea hydrothermal white smoker

Methanocaldococcus vulcanius

Irregular cocci

49–89

80

H2 only

Deep-sea hydrothermal chimney

Methanotorris igneus

Irregular cocci

45–91

88

H2 only

Coastal hydrothermal system

Methanotorris formicicus

Irregular cocci

55–83

75

H2 only

Deep sea black smoker chimney

Methanomicrobiales Rods

35–55

50

H2, formate

Methanogenic sewage sludge

Irregular cocci

37–65

55–60

H2, formate

High temperature effluent channel

Methanogenium frittonii Irregular cocci

26–62

57

H2, formate

Nonthermal freshwater sediments

Methanolinea tarda Methanomicrobiaceae Methanoculleus thermophilus

Methanosarcinales Methanosarcinaceae Methanosarcina thermophila

Sarcina

35–55

50

H2 , methylamines, methanol, acetate

Sludge digestor

Methanohalobium evestigatum

Polygonal cocci

30–60

50

Tri- di-mono methylamines

Saline reservoirs in the Crimea

Methanomethylovorans thermophila

Irregular cocci

42–58

50

Methanol, methylated amines

UASB bioreactor

Rods

37–70

60

Acetate

Thermophilic anaerobic bioreactor

Small cocci

50–70

65

Methanol, methylated amines

Oil-production water

Methanosaetaceae Methanosaeta thermophila Methermicoccaceae Methermicoccus shengliensis

54

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

. Table 1 (Continued) Temperature Range Morphology ( C)

Species

Optimum ( C)

Methanogenesis substrates Typical habitats

Methanopyrales Methanopyraceae Methanopyrus kandleri

Rods

84– 110

98

H2 only

Hydrothermally heated deep sea

into methane (Methanohalobium, Methanomethylovorans, and Methermicoccus species). One species is considered to be the only thermophilic strictly acetoclastic methanogen (Methanosaeta thermophila) known to date; a single species (Methanosarcina thermophila) can use acetate, methanol, and methylated amines together with hydrogen (> Table 1).

2

Importance of Thermophilic Methanoarchaea in the Breakdown of Organic Matter

As already stated, thermophilic methanoarchaea occupy the terminal position in the anaerobic breakdown of organic matter. This is the case in anaerobic digesters operating at high temperature, where they are essential for the mineralization of biopolymers, converting hydrogen and acetate produced by fermentative or syntrophic bacteria into methane. These last microorganisms require a microbial association with a hydrogenotrophic partner (e.g., methanogen) to oxidize fatty acids (e.g., Thermosyntropha). This feature was termed ‘‘interspecies hydrogen transfer’’ when first discovered in a mesophilic methanogenic coculture (Garcia et al., 2000). Nonobligate interspecies hydrogen transfer has also been demonstrated with defined cocultures of thermophilic hydrogenotrophic methanogens with fermentative bacteria (Garcia et al., 2000). In the absence of organic matter, thermophilic methanogenesis occurs mainly from geochemical hydrogen formed as part of the geological process. This is the case in hot springs, deep-sea hydrothermal vents and in subsurface petroleum reservoirs, where the contribution of mesophilic hydrogenotrophic methanoarchaea to crude-oil degradation has recently been established (Jones et al., 2008). A similar participation of thermophilic hydrogenotrophic methanoarchaea in this anaerobic biological process may thus be hypothesized in deep oil reservoirs. Methanoarchaea can play a major ecological role (through geomicrobiological processes) in hot springs as primary producers owing to their ability to oxidize hydrogen and reduce CO2. However, the methanoarchaeal diversity occurring in these hot niches reduces to Methanothermobacter and Methanothermus species that readily grow on hydrogen. Interestingly, Methanothermus fervidus and M. sociabilis, the only representatives of the genus Methanothermus, have been isolated only in solfatara fields in southwest Iceland (> Table 1), and are believed to grow endemically in that country. Compared with hot springs, much more information has been obtained for methanoarchaea inhabiting oil reservoirs (Jeanthon et al., 2005; Magot et al., 2000) and deep-sea hydrothermal vents (Jeanthon, 2000; Miroshnichenko and Bonch-Osmolovskaya, 2006). Here, we focus on methanoarchaea inhabiting these two deep extreme environments.

685

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54

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

. Figure 1 Neighbor-joining phylogenetic tree, based on 16S rRNA gene sequences, showing the relationships between thermophilic (boldface type) and mesophilic methanoarchaea (only the type species of each mesophilic genus is given).

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

3

Methanoarchaea from Extreme Environments

3.1

Oilfield Reservoirs

54

Among subterrestrial ecosystems, oilfields represent unusual anaerobic environments because of the diversity of their physical, chemical, and geochemical conditions. In these subterrestrial ecosystems, temperature increases by 2–3 C per 100 m depth (temperatures within the oil reservoirs may range from 30 to 180 C) and geological formations can differ drastically from one oilfield to another (saline to hypersaline formations). Despite the extreme physical and chemical conditions prevailing in oil reservoirs, the existence of microbial life has now been clearly established in these ecosystems and a complex and physiologically diverse microbial assemblage of mesophilic to hyperthermophilic and/or halophilic to hyperhalophilic anaerobic microorganisms distributed among the domains Bacteria and Archaea has already been reported (Magot et al., 2000; Ollivier and Cayol, 2005). Microbes originating from oilfield facilities include fermentative, sulfate-reducing, and methanogenic microorganisms. Methanogenesis has been reported several times in deep oil-bearing strata (Magot et al., 2000) and thermophilic methanoarchaea have thus been isolated from hot oil reservoirs. They include members of the Methanobacteriaceae, Methanococcaceae, Methanosarcinaceae, and Methermicoccaceae families. Representatives of the Methanobacteriaceae family inhabiting oilfield ecosystems are rod-shaped microorganisms of the genus Methanothermobacter. This is the case for M. thermautotrophicus, and phenotypic variants of ‘‘M. thermaggregans (formely Methanobacterium thermaggregans)’’ recovered from hot oil reservoir fluids (Magot et al., 2000). The isolation of this last archaeon from the high temperature and moderately saline San Miguelito oilfield provides evidence that virgin oil reservoirs can contain an in situ methanogenic population. Methanothermococcus thermolithotrophicus was isolated from a North Sea oilfield reservoir (Magot et al., 2000). Close relatives of M. thermolithotrophicus have also been identified from 16S rDNA gene libraries in the Troll formation in the North Sea (Dahle et al., 2008). Using the same culture-independent techniques, representatives of the genus Methanothermococcus were revealed from samples collected at Samotlor high-temperature oil reservoirs (Western Siberia, Russia). The archaeal libraries (138 clones) were mostly composed of microorganisms belonging to sulfate-reducers and methanogens, the latter microorganisms being represented by an important group of sequences clustering with Methanocaldococcus infernus (97% similarity) (Jeanthon et al., 2005), a methanoarchaeon to date only isolated from deep-sea hydrothermal vents in the Mid-Atlantic Ridge (Jeanthon et al., 1998). We note that both Methanothermobacter and Methanothermococcus species share a common feature in their use of hydrogen, which is most probably their primary source of energy in oilfields. Hydrogen generation in oil reservoirs may be the result of (i) abiotic reactions at high temperature in the deep biosphere or (ii) biotic reactions via fermentation processes or hydrocarbon oxidation. It has been recently demonstrated in heavily degraded oils that methanogenic hydrocarbon degradation occurs predominantly through syntrophic oxidation of alkanes to acetate and hydrogen (Jones et al., 2008). Experiments conducted in North Sea oil reservoirs have shown that methane is produced in enrichment cultures with acetate as the substrate at 60, 80 and 92 C and that acetate can accumulate to concentrations of up to 20 mM in some of these reservoirs (Magot et al., 2000). Methanogenesis from acetate-enriched media has also been observed in high-temperature formations in the Mykhpai field (western Siberia) (Magot et al., 2000). Besides culturedependent techniques, molecular phylogenetic surveys of high-temperature sulfur-rich

687

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54

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

oil reservoirs in California based on the analysis of sequences of genes encoding for the 16S rRNA (Orphan et al., 2000) revealed the presence of seven clones (out of 154) that were distantly related ( Table 1). The only thermophilic methylotrophic methanoarchaeon known to grow at temperatures over 60 C was recently isolated from the Shengli oilfield in China and named Methermicoccus shengliensis (> Table 1). The role of these methylotrophs in oilfield waters is poorly understood as their metabolism cannot be linked to hydrogen and/or hydrocarbon oxidation, considered as the two major energy sources available in situ.

3.2

Deep-Sea Hydrothermal Vents

Since the discovery of deep-sea hydrothermal vents in the eighties in the Galapagos Rift, much attention has been paid to the study of thermophilic/hyperthermophilic anaerobic and microaerobic bacterial and archaeal communities inhabiting these deep hot ecosystems (Jeanthon, 2000; Miroshnichenko and Bonch-Osmolovskaya, 2006). Despite the drastic physical and chemical conditions prevailing in situ, which include high temperature and hydrostatic pressure together with elevated concentrations of heavy metals discharged by the hydrothermal fluid of black smokers at temperatures in the range 300–400 C into the low temperature (approx. 4 C) oxygenated seawater, there is evidence that numerous chemolithotrophic and chemoorganotrophic microorganisms (domains Bacteria and Archaea) inhabit this extreme environment. Within the domain Archaea, several thermophilic methanoarchaea of the order Methanococcales or Methanopyrales have been recovered from deep-sea hydrothermal vents (Miroshnichenko, 2004; Miroshnichenko and Bonch-Osmolovskaya, 2006). Methanopyrus kandleri, the only representative of this genus in the order Methanopyrales is characterized by the highest growth temperature ever recorded for any methanoarchaea

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

54

known to date (optimum 98 C, maximum 110 C). As hydrogen produced by abiotic reactions at high temperature between basaltic rock and seawater is present at high concentrations in the dissolved gases of hydrothermal fluids, hydrogenotrophic methanoarchaea may play significant ecological and geomicrobiological roles, notably in the carbon cycle of these deep hot environments and also possibly in the sulfur cycle, as some methanoarchaea have been reported as potential sulfur reducers. We note that most of these methanoarchaea (e.g., members of the Methanocaldococcaceae and Methanopyraceae families) isolated to date from such environments use hydrogen as their sole energy source and may grow autrophically, suggesting that they may participate actively in the primary production of organic matter together with the other lithotrophic anaerobic or microaerobic microbes thriving in the deep hot ocean environment (e.g., Desulfurobacterium, Persephonella). Of interest is the observed endemic character of Methanocaldococcus species, and also of Methanopyrus kandleri, in particular in the deep-sea hydrothermal vents, suggesting that they most probably should be considered as indigenous inhabitants of these extreme environments (Miroshnichenko and Bonch-Osmolovskaya, 2006). Besides elevated temperatures, methanoarchaea also have to withstand hydrostatic pressure, and for example, Methanocaldococcus jannaschii was found to be barophilic. As already indicated, hydrogenotrophic methanoarchaea isolated from deep-sea hydrothermal vents are essentially represented by members of the genera Methanocaldococcus, Methanothermococcus, and Methanotorris, order Methanococcales and Methanopyrus, order Methanopyrales (> Table 1, > Fig. 1). The ecological significance of members of these two orders within the deep hot marine environment has also been revealed by 16S rRNA and mcrA (methyl coenzyme M reductase) gene sequence analysis (Dhillon et al., 2005; Miroshnichenko and Bonch-Osmolovskaya, 2006). Interestingly, five mcrA clones retrieved from a deep-sea hot environment (Guaymas Basin) were found to have Methanoculleus thermophilus, a thermophilic methanogen growing optimally at 55–60 C, as their closest relative, suggesting that a wide range of thermophilic methanoarchaea (from thermophiles to hyperthermophiles) may thrive in the deep geothermally heated marine sediments (Dhillon et al., 2005). By contrast, no work is reported on the isolation of thermophilic acetoclastic methanoarchaea, which may contribute to the complete anaerobic oxidation of organic matter in deep-sea hydrothermal vents. However, the 16S rRNA clone library from homogenized Guaymas Basin sediments indicated some phylotypes closely related to cultured acetoclastic thermophilic species (e.g., members of the order Methanosarcinales) and to clones from hot petroleum reservoirs (Dhillon et al., 2005). In addition, radioisotopic measurements for deep-sea hydrothermal vents of the East Pacific Rise provided evidence of methane production from acetate at 80 C (Miroshnichenko, 2004). This temperature is the highest recorded to date for acetoclastic methanogenesis. All these results thus suggest that thermophilic acetoclastic methanogenesis may be expected in deep hot marine environments. If not, like in oilfield ecosystems, in situ activity of syntrophic acetoclastic microorganisms associated with hydrogenotrophic methanoarchaea may be hypothesized. Also, acetoclastic iron-reducers (Geoglobus ahangarii, Deferribacter abyssi) and nitrate-reducers (Caldithrix abyssi) have been isolated from such environments (Miroshnichenko and Bonch-Osmolovskaya, 2006).

4

Conclusion

Thermophilic methanoarchaea are known to thrive in hot made-man environments (e.g., compost and high-temperature digestors), where they contribute significantly through

689

690

54

Thermophilic Methanoarchaea Inhabiting Hot Ecosystems

hydrogen and acetate utilization to anaerobic oxidation of organic matter. They are also found in extreme niches of the planet, both terrestrial, e.g., hot springs, and subterrestrial, such as oilfield environments and deep-sea hydrothermal vents. In these two deep environments, where they have had to adapt to drastic physical and chemical conditions, including temperature, pressure and salinity, the primary source of energy is hydrogen. This gas can be produced from high-temperature abiotic reactions or fermentation processes. Although thermophilic acetoclastic methanogens have been identified in these subterrestrial environments by analyzing 16S rRNA gene clone libraries (Dhillon et al., 2005; Jeanthon, 2000; Jeanthon et al., 2005; Magot et al., 2000), none of these microorganisms has been isolated to date. Syntrophic acetate degradation may also be a significant hydrogen source for hydrogenotrophic methanoarchaea, notably in oil reservoirs. By contrast, in deep-sea hot environments, even though the existence of syntrophic acetate degradation cannot be totally ruled out, there is evidence that anaerobic acetate oxidation can be achieved through iron- or nitrate reduction processes, thus making complete anaerobic organic matter oxidation easier in this ecosystem through the high availability of mineral electron acceptors in situ. It is of note that hydrogenotrophic methanoarchaea originating from oilfield environments comprise only thermophiles with an optimum temperature for growth of 60–65 C, whereas most of those originating from deep-sea hydrothermal vents are recognized as hyperthermophiles, growing optimally at temperatures higher than 80 C and as endemic to the deep hot ocean. This is not the case for hydrogenotrophic oilfield isolates, which have been found in other hot habitats (e.g., high temperature digestors). Only one thermophilic strictly methylotrophic methanoarchaea species growing optimally at 65 C (Methermicoccus shengliensis) has been isolated to date from oil reservoirs, but its ecological role in the deep hot reservoirs has not been elucidated. However, the role of thermophilic hydrogenotrophic methanoarchaea should be considered to be of ecological significance in the deep hot ecosystems where they can be regarded as active primary producers (deep-sea and oilfield environments) and possibly important contributers to crude-oil biodegradation in deep petroleum reservoirs, since the importance of Methanoarchaea in this biodegradation has been recently demonstrated in shallow low temperature reservoirs (Jones et al., 2008). These features make them essential actors in the overall biogeochemistry of the carbon cycle in the deep biosphere.

5

Research Needs

Although acetoclastic methanogenesis has been evidenced through molecular studies and activity measurements in the deep hot biosphere, isolation and characterization of methanoarchaea with the ability to oxidize acetate still remain unfruitful and should be considered as a challenge in future for microbiologists. This is true in deep sea hydrothermal vents, but also in deep oil reservoirs. In the latter ecosystems, the contribution of hydrogenotrophic methanoarchaea in crude oil biodegradation has been only demonstrated in shallow lowtemperature reservoirs, but still needs to be elucidated in deep high-temperature reservoirs. Finally, further microbiological investigations in the deep hot biosphere should help in possibly extending the upper limit temperature for the growth of methanoarchaea.

Acknowledgments Many thanks to Pierre Roger for revising the manuscript.

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References Dahle H, Garshol F, Madsen M, Birkeland N-K (2008) Microbial community structure analysis of produced water from a high-temperature North Sea oil-field. Anton Leeuwenhoek 93: 37–49. Dhillon A, Lever M, Lloyd KG, Albert DB, Sogin ML, Teske A (2005) Methanogen diversity evidenced by molecular characterization of methyl coenzyme M reductase A (mcrA) genes in hydrothermal sediments of the Guaymas Basin. Appl Environ Microbiol 71: 4592–4601. Garcia JL, Patel BKC, Ollivier B (2000) Taxonomic, phylogenetic, and ecological diversity of methanogenic Archaea. Anaerobe 6: 205–226. Jeanthon C (2000) Molecular ecology of hydrothermal vent microbial communities. Anton Leeuwenhoek 77: 117–133. Jeanthon C, L’Haridon S, Reysenbach AL, Vernet M, Mesner P, Sleytr UB, Prieur D (1998) Methanococcus infernus sp. nov., a novel hyperthermophilic lithotrophic methanogen isolated from a deep-sea hydrothermal vent. Int J Syst Bacteriol 48: 913–919. Jeanthon C, Nercessian O, Corre E, Grabowski-Lux A (2005) Hyperthermophilic and methanogenic Archaea in oil fields. In Petroleum Microbiology. B Ollivier and M Magot (eds.). Washington, DC: ASM Press, pp. 55–69. Jones DM, Head IM, Gray ND, Adams JJ, Rowan AK, Aitken CM, Benett B, Huang H, Brown A, Bowler BFJ, Oldenburg T, Erdmann M, Larter SR (2008) Crude-oil biodegradation via methanogenesis in

subsurface petroleum reservoirs. Nature 451: 176–180. Liu Y, Whitman WB (2008) Metabolic, phylogenetic, and ecological diversity of the methanogenic Archaea. Ann N Y Acad Sci 1125: 171–189. Magot M, Ollivier B, Patel BKC (2000) Microbiology of petroleum reservoirs. Anton Leeuwenhoek 77: 103–116. Miroshnichenko ML (2004) Thermophilic microbial communities of deep-sea hydrothermal vents. Microbiology 73: 1–13. Miroshnichenko ML, Bonch-Osmolovskaya EA (2006) Recent developments in the thermophilic microbiology of deep-sea hydrothermal vents. Extremophiles 10: 85–96. Ollivier B, Cayol JL (2005) The fermentative, ironreducing, and nitrate-reducing microorganisms. In Petroleum Microbiology. B Ollivier and M Magot (eds.). Washington, DC: ASM Press, pp. 71–88. Orphan VJ, Goffredi SK, Delong EF, Boles JR (2003) Geochemical influence on diversity and microbial processes in high temperature oil reservoirs. Geomicrobiol J 20: 295–311. Orphan VJ, Taylor LT, Hafenbradl D, Delong EF (2000) Culture-dependent and culture-independent characterization of microbial assemblages associated with high temperature petroleum reservoirs. Appl Environ Microbiol 66: 700–711. Wagner ID, Wiegel J (2008) Diversity of thermophilic anaerobes. Ann N Y Acad Sci 1125: 1–43.

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55 Mammalian Digestive Tract G. N. Jarvis1,* . D. Al-Halbouni2 1 New Zealand Trade & Enterprise, Biotechnology & Agritechnology Sector, Wellington, New Zealand 2 Institute for Biology, Unit of Soil Ecology, RWTH Aachen University, Aachen, Germany *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 694

2 Methanogenic Archaea Communities in Humans . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 695 2.1 Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 695 2.2 Composition and Taxonomy of Methanogenic Populations in the Human Colon . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 695 2.3 Ecophysiology and Genetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 697 2.4 Microbial Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 698 2.5 Methanogenic Archaeal Communities in Other Monogastric Ecosystems . . . . . . . . . . 698 3 3.1 3.2 3.3 3.4

Methanogenic Archaeal Communities in Ruminants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 698 Overview . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 698 Composition and Taxonomy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 699 Ecophysiology and Genetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 700 Microbial Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 700

4 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 701 4.1 Human Methanogenic Archaea and Disease . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 701 4.2 Methanogenic Community Structure, Succession and Methane Mitigation . . . . . . . . 702 5

Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 702

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_55, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Methanogenic archaea associated with mammalian digestive tracts have a limited metabolic range, yet in some cases have significant phylogenetic diversity. Interest in this group of archaea is increasing as a result of their role in production of biological methane – a critical greenhouse gas. It is clear that representatives from the genus Methanobrevibacter predominate in a number of mammalian gastrointestinal ecosystems. However, the total archaeal species composition, methanogen ecophysiology and the methanogen–microbe interactions does indicate a degree of diversity in these ecosystems. Further studies will help resolve how significant the species diversity is as regards the methanogenic archaeal community structure, and will further aid our understanding of the interactions with both the host and other microbes within mammalian gastrointestinal ecosystems.

1

Introduction

Biological methane production has in recent times attracted significant public interest because of its critical role as a major green-house gas. According to Liu and Whitman (2008), the current total global methane emissions are 500–600 Tg CH4 y 1, with the majority, approximately 74%, being derived from biological methanogenesis. Methanogens are members of the domain Archaea and fall within the kingdom Euryarchaeota. They are obligate anaerobes and require both anoxic and highly reduced environmental conditions i.e., redox potential below –330 mV for growth (Sowers, 1995). Methanogens all share one metabolic characteristic that differentiates them from other prokaryotic forms of life – the ability to utilize carbon dioxide or methyl groups as the terminal electron acceptor and yield methane as the major catabolic product from metabolic processes (Liu and Whitman, 2008). Despite methanogens having both a comparatively narrow range of substrates and a requirement for fastidiously anaerobic, reduced environmental conditions for growth, they are relatively ubiquitous and are found in a range of highly diverse environments and ecosystems, including those associated with: organic matter decomposition e.g., peat bogs, mammalian and insect gastrointestinal tracts; geochemical activities e.g., deep sea hydrothermal vents, and; anthropogenic activities e.g., anaerobic digesters (Liu and Whitman, 2008; Sowers, 1995). While the natural production of combustible gases (methane) was described first during Roman times, it was not until 1776 that the Italian physicist Alessandro Volta made an association between ‘‘combustible air’’ and the presence of plant material in aqueous sediments. The involvement of microbes in methanogenic processes was first reported in 1868 by Bechamp (Sowers, 1995). Surprisingly, significant research and confirmed isolations and association of methanogens with the gastrointestinal tract of mammals has only occurred in the latter part of the twentieth century with pioneering researchers using classic isolation studies on microbial populations associated with the rumen ecosystem (Paynter and Hungate, 1968) and human colonic ecosystem (Nottingham and Hungate, 1968). Therefore, in comparative historical terms, mammalian gastrointestinal ecosystems represent some of the more recently studied ecosystems as regards biological methanogenesis. However, the combination of enhanced classic cultivation techniques such as the Hungate technique (Hungate, 1966), and modern molecular biology techniques, have enabled us to

Mammalian Digestive Tract

55

gain insight about methanogens associated with these gastrointestinal ecosystems in a more rapid manner.

2

Methanogenic Archaea Communities in Humans

2.1

Overview

The question arises as to whether mammalian gastrointestinal ecosystems could be defined as high-lipid content ecosystems. In the case of humans there is no doubt that high dietary lipid intake occurs as in the case of Western diets where up to 40% of total dietary energy is derived from fats – mainly triglycerides (Cordain et al., 2005). However as the majority of mammalian lipid metabolic processes occur in small intestine, then the colon itself has little, if any, exposure to dietary lipids. To date, a limited number of methanogens have been found associated with only three sites within humans: the oral cavity (Methanobrevibacter oralis), vagina (Mbb. smithii) and distal gut (Mbb. smithii and Methanosphaera stadtmanae) (Lepp et al., 2004). While the total microbial community in the human gut numbers between 10 and 100 trillion cells, the largest numbers are associated with the distal gut (colon) (Gill et al., 2006). Within the colon fermentation of organic substrates by anaerobic microbial communities, generally dominated by bacteria from the division Bacteroidetes and Firmicutes, leads to production of short chain fatty acids and CO2 (Gill et al., 2006; Liu and Whitman, 2008). Disposal of reducing equivalent is achieved via the production of H2, which is maintained at a low partial pressure in the colon via its removal by hydrogenotrophic microbes including acetogens, sulfatereducing bacteria and methanogens (Liu and Whitman, 2008). Between 34 and 50% of healthy humans harbor methanogens in their colons based on studies analyzing breath methane levels (Levitt et al., 2006; Pochart et al., 1992). Interestingly, most colonic methane is excreted in flatus from humans, and only small proportions are either absorbed into the bloodstream or released in the breath. The total number of colon-associated methanogens resident in methane-producing humans is between 108 and 1010 per gram feces compared to < 102–5.0  106/g feces in non-methane producing humans (Liu and Whitman, 2008; Scanlan et al., 2008).

2.2

Composition and Taxonomy of Methanogenic Populations in the Human Colon

The first isolation and identification of human methanogens was achieved in 1968 and yielded identical isolates denoted as strains of ‘‘Methanobacterium ruminantium’’ from the feces of four individuals (Nottingham and Hungate, 1968). However, this epithet was later replaced and its members reclassified into two species – Methanobrevibacter smithii and Methanobrevibacter ruminantium (Miller et al., 1982). Despite the large numbers of methanogens often associated with this ecosystem, to date, only two species (> Table 1) have been formally isolated and identified from the human colon – Methanobrevibacter smithii and Methanosphaera stadtmanae (Miller and

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. Table 1 Representative methanogenic species known to be associated with human and animal gastrointestinal tracts Optimum growth Genus species

Morphology

Substrates1

pH

Temp (C)

Cofactors for growtha

Hosts

Methanobrevibacter Mbb. smithii

Coccobacillus

H, F

6.9–7.4

37–39

Ac, B vits

Mbb. gottschalkii

Coccobacillus

H

7.0

37

Ac, Trp, YE Horse, pig,

Mbb. thaueri

Coccobacillus

H

7.0

37

Ac, Trp, YE Bovinea

Mbb. woesei

Coccobacillus

H, F

7.0

37

Ac, Trp, YE Goose

Mbb. wolinii

Coccobacillus

H,

7.0

37

Ac, Trp, YE, CoM, 2-MB

Sheepa

Mbb. ruminantium

Coccobacillus

H, F

6.3–6.8

37–39

Ac, AA, CoM, 2MB

Cattleb

Mbb. millerae

Coccobacillus

H, F

7.0–8.0

36–42

Ac, YE, Trp Cattleb

Mbb. olleyae

Coccobacillus

H, F

7.5

36–40

Ac

Sheepb

Mbb. oralis

Coccobacillus

H

6.9–7.4

36–38

VFA, FE

Human

Mb. formicium

Rod

H, F, 2-P, 2-B

6.6–7.8

37–45

None

Cattleb

Mb. bryantii

Rod

H, F, 2-P, 2-B

6.9–7.2

37–39

None

sheepb

6.5–7.5

30–40

None

Cattleb

Human, cattleb

Methanobacterium

Methanosarcina Ms. barkeri

Pseudosarcina Ac, Me, H2

Sheepb Methanosphaera Msp. stadtmanae

Coccus

H, Me

6.5–6.9

36–40

Ac

Human bovineb Ovineb

Methanoculleus Mc. olentangyi

Irreg. coccus

H, F

6.5–7.0

37

None

Deerb

Curved rod

H, F

6.1–6.9

40

RF

Cattleb

Methanomicrobium Mm. mobile

Sheepb Substrates: Ac acetate; H H2/CO2; F formate; Me methanol; 2-P propan-2-ol; 2-B butan-2-ol. Co-factors: Ac acetate; B-vits B vitamin mix; trp trypticase; YE yeast extract; AA amino acid mix; CoM coenzyme M; 2-MB 2 methyl butyrate; RF rumen fluid; VFA volatile fatty acid mix; FE fecal extract a Isolated from feces/colon b Isolated from rumen

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Wolin, 1985; Miller et al., 1982). Mbb. smithii is the predominant methanogen associated with this ecosystem and has been shown to compose up to 10% of the total anaerobic population in the colon (Belay et al., 1990; Miller and Wolin, 1986). However, a recent culture-independent study has also indicated the potential presence of Methanobrevibacter oralis-like and Methanosarcina-like species in the colon (Scanlan et al., 2008) (See > Chapter 42, Vol. 1, Part 7).

2.3

Ecophysiology and Genetics

Methane in the human colon is mainly produced from H2 and CO2, and there is little, if any, aceticlastic methanogenesis. This is due to the short residence time in the gut and slower growth rates of the methanogens using acetate as a substrate (Liu and Whitman, 2008). Because Mbb. smithii is capable of using not only H2 and CO2, but also bacterial fermentation products such as formate, sometimes albeit weakly (Miller et al., 1982) for growth, then it seems it may be well adapted to its role as the predominant methanogen in the human gastrointestinal system. Genomic analysis indicated that Mbb. smithii has the ability to up-regulate the production of the formate utilization gene cluster – formate transporter FdhC and formate dehydrogenase subunits FdhAB – which may explain the variable growth phenotype observed with formate (Miller et al., 1982; Samuel et al., 2007). Assimilation of acetate rather than use as a growth substrate, is explained by the presence of an ‘‘incomplete tricarboxylic acid cycle’’ in the Mbb. smithii proteome (Samuel et al., 2007). Further molecular analysis of the Mbb. smithii genome indicated that it codes for enzymes involved in the synthesis of human gut mucosal – like surface glycans and adhesion-like proteins (Samuel et al., 2007) There is also some genomic evidence that Mbb. smithii may be able to utilize ethanol and methanol – both of which are bacterial fermentation products found in the colon ecosystem (Liu and Whitman, 2008; Samuel et al., 2007; Scanlan et al., 2008). In contrast Msp. stadtmanae has a much more limited growth substrate range, as it can only reduce methanol with H2 (Fricke et al., 2006; Liu and Whitman, 2008; Scanlan et al., 2008). Energy conservation is accomplished by coupling of ADP phosphorylation with methanol reduction using H2 to methane. Both energy metabolism and conservation processes are achieved by five enzyme complexes: methanol:coenzyme M methyltransferase, methylcoenzyme M reductase, heterodisulfide reductase, non-F420 reducing hydrogenase, and proton-translocating ATPase. The limited substrate range of Msp. stadtmanae is also due to it lacking protein coding sequences (CDS) including those for synthesis of carbon monoxide dehydrogenase/acetyl coenzyme A complex and molybdopterin – which are required for reduction of CO2 to methane or oxidation of methanol to CO2. Interestingly, those CDS are present in all other sequenced methanogens (Fricke et al., 2006). Lack of the CDS for the nickel-containing carbon monoxide dehydrogenase/acetyl CoA decarbonylase complex also explains why Msp. stadtmanae cannot use acetate as an energy substrate, and is instead dependent on it as a carbon source for growth (Fricke et al., 2006). Furthermore, the lack of genes for the 2-oxoisovalerate: ferredoxin oxidoreductase complex, which is required for assimilation of branched chain fatty acids also explained why this methanogen does not require branched chain fatty acids as a co-factor for growth. Interestingly, as with Mbb. smithii, the genome of Msp. stadtmanae also encodes for both human-like surface glycans and adhesions, which lead to speculation that these specific characteristics were part of

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adaptation to host environment – the human large intestine (Fricke et al., 2006; Samuel et al., 2007). Despite this latter point, it is tempting to postulate that the limited metabolic range may explain why Msp. stadtmanae is generally found in a much lower incidence in this ecosystem compared to Methanobrevibacter isolates (Gill et al., 2006; Lin and Miller, 1998; Scanlan et al., 2008), although more studies will be required to confirm this observation (See > Chapter 36, Vol. 1, Part 6).

2.4

Microbial Interactions

Removal of H2 serves as an important function in gastrointestinal systems, and it is one that methanogens clearly excel at. It has been observed that methanogenic removal of H2 improves efficiency of polysaccharide metabolism, short chain fatty acid production and enhanced bacterial growth. This is further supported by the human distal gut microbiome yielding methanogenic gene clusters essential for these functions (Gill et al., 2006). Methanogenic archaea, acetogenic and sulfate reducing bacteria (SRB) all exist within the human colon ecosystem. In the case of the SRB and methanogens it appears that this interaction is a case of co-existence rather than competition (Pochart et al., 1992; Strocchi et al., 1994). Obviously this is surprising given that under standard conditions methanogens should be out-competed by SRB (Liu and Whitman, 2008). In low or non-methane producing humans the number of acetogenic bacteria predominate and may out-compete the methanogens (Bernalier et al., 1996). While negative correlations between methanogenesis and bowel movements, and fecal butyrate levels have been recorded (Liu and Whitman, 2008), the exact drivers for the abundance or absence of methanogens in the human colon ecosystem have yet to be fully elucidated.

2.5

Methanogenic Archaeal Communities in Other Monogastric Ecosystems

Studies on other monogastric animals have included a variety of species including the pig, horse, monkey, baboon, rhinoceros, giant panda, turkey and goose, hippopotamus, and chickens (Jensen, 1996; Lin and Miller, 1998; Miller and Lin, 2002). Examples of the species from these ecosystems are included elsewhere (> Table 1) but in all cases the predominant genus present in the methanogenic communities in these monogastric gastrointestinal tracts is Methanobrevibacter. However, it is interesting to note that both the turkey and chicken gastrointestinal ecosystems also harbor species belonging to the genus Methanogenium (Jensen, 1996).

3

Methanogenic Archaeal Communities in Ruminants

3.1

Overview

The rumen is the main site of microbial fermentation of plant material in wild and domesticated livestock and ruminant mammals, and is home to a multitude of microorganisms.

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Indeed on a per milliliter (ml.) basis this ecosystem harbors: 1010–1011 bacteria, 104–106 ciliate protozoa, 103–105 zoospores (anaerobic fungi), 108–109 bacteriophages and 108–1010 methanogenic archaea. It is estimated that approximately 3–4% of total ruminal microbes are methanogens (Janssen and Kirs, 2008; Kamra, 2005; Liu and Whitman, 2008; Morvan et al., 1996). Ruminal methanogenesis has in recent times attracted a lot of scientific and public interest. This is mainly due to the significant role that this ecosystem has as a major biologic methanogenesis source, which is estimated to account for 13–19% of global methane emissions (Liu and Whitman, 2008). Ruminant diets, depending on animal husbandry practices, may contain between 3–9% lipid on a dry matter basis – but generally the diet does not contain more than 6% fat. This is to avoid adverse effects on rumen digestion processes and animal nutritive health (Jenkins, 1993). Assuming a standard pasture-based diet (fed ad libitum, 4% lipid content) then we can conservatively estimate that that lipid intake, in the form of plant galactolipid, phospholipids and sulfolipids, by an individual bovine and ovine will be approximately 210 kg lipid/per year and 25 kg lipid/per year respectively per animal (Ulyatt et al., 2002). Assuming similar dry matter intake values and ad libitum access, then for concentrate or cereal-based diets it is conceivable that an individual bovine animal and its rumen microbial ecosystem could be exposed to over 600 kg lipid/per year mainly in the form of triglycerides as the major lipid. However, despite the data outlined above it seems unlikely that one could, senso stricto, define this as a high lipid content ecosystem. Interestingly, the use of lipids to modify/reduce ruminal methanogenesis has received significant attention recently, given the current interest in mitigating greenhouse gas emissions from livestock (Calsamiglia et al., 2007).

3.2

Composition and Taxonomy

The composition of the ruminal methanogenic population is obfuscated by the fact that the rumen is not a homogenous ecosystem, and methanogens are associated with three distinct phases, namely: the rumen fluid phase; the feed particle phase and; in symbiotic relationships with protozoa (Sharp et al., 1998; Shin et al., 2004; Whitford et al., 2001). Nevertheless, a number of studies have indicated that members of the genera Methanobrevibacter and Methanomicrobium are the predominant ruminal methanogens (> Table 1) irrespective of animal species or spatial location in the rumen ecosystem (Shin et al., 2004; Skillman et al., 2004; Tatsuoka et al., 2004; Whitford et al., 2001; Wright et al., 2004; Yanagita et al., 2000). In a recent meta-analysis of clone library sequence data it was established that a 3rd genus-level group termed RCC (rumen cluster C) of uncultured archaea with significant sequence variation exists in the rumen ecosystem (Janssen and Kirs, 2008). Rather surprisingly, there have only been a limited number of culture dependent studies on the rumen ecosystem. These studies have yielded isolates belonging to the eight species Methanobacterium formicium, Mb. bryantii, Methanobrevibacter ruminantium, Mbb. millerae, Mbb. olleyae, Methanomicrobium mobile, Methanoculleus olentangyi, and Methanosarcina barkeri (Jarvis et al., 2000; Oppermann et al., 1957; Paynter and Hungate, 1968; Rea et al., 2007; Skillman et al., 2004; Smith and Hungate, 1958). Interestingly, while phylogenetic studies highlight the presence of members of the genus Methanimicrococcus in rumen samples – no pure culture has been isolated from the rumen to date (Janssen and Kirs, 2008).

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Overall, numerous studies indicate the exact genus and species composition and abundance seems to vary in the rumen ecosystem. This finding is hardly surprising given the vast array of potential physical niches, and interactions with other rumen microbes. As a result, Janssen and Kirs (2008) noted that co-existence rather than competition is likely to occur between different methanogens in the rumen ecosystem. In contrast to the rumen ecosystem, less data is available on the colon ecosystem of ruminants. The data available has however indicated that the predominant genus in this ecosystem is Methanobrevibacter (Lin and Miller, 1998; Liu and Whitman, 2008).

3.3

Ecophysiology and Genetics

While H2/CO2 is the major substrate of rumen methanogens, other potential substrates including formate, acetate, methylamine (from the degradation of choline) and methanol (from the degradation of plant polymers) do occur in the rumen (Stewart et al., 1997). Based on both enumeration and phylogenetic analysis it is clear that the majority of the methanogenic archaea associated with the rumen are predominantly hydrogen-utilizing (Janssen and Kirs, 2008; Liu and Whitman, 2008; Morvan et al., 1996; Wolin et al., 1997). Indeed, while formate has been identified as a substrate for methanogens, its half life in the rumen is very short and it is unlikely to ever have a major role in ruminal methanogenesis (Hungate, 1966). However, there is less clear evidence for any role for acetate as a methanogenic substrate in the rumen ecosystem, especially given its rapid absorption across the rumen epithelium and utilization by the host (Jenkins, 1993). Given the increased interest in rumen methanogenesis it is rather surprising that no publicly available genome sequence has been produced for a rumen methanogen (Liu and Whitman, 2008). A private consortium has recently reported it has fully sequenced the genome of Methanobreviobacter ruminantium yielding a genome size of approximately 3 mega-base pairs (Mbp) and harboring over 2,220 genes with approximately half of these having no known function. Furthermore, all genes associated with methanogenesis have been found in this archaeal species and are conserved when compared to other Methanobacteriales genome sequences (Attwood et al., 2008).

3.4

Microbial Interactions

One of the most critical interactions between methanogenic archaea and other members of the rumen ecosystem revolves around interspecies H2 transfer. It is of some note that the estimated 800 l of H2 produced by ruminal fermentation processes is converted into 200 l of methane by a 500 kg cow, with only a minor trace of the H2 being found in the gaseous phase of the rumen (Wolin et al., 1997). Maintenance of a low H2 partial pressure by methanogens has a significant impact on cellulose fermentation as well as end product profiles, ATP yields and growth rates by hydrolytic fermentative bacteria in the rumen (Kamra, 2005; Wolin et al., 1997). These syntrophic relationships however do not lead to a complete benefit for the host, as methanogenesis has been shown to be responsible for a net loss of 6–12% of ingested feed energy as methane (Liu and Whitman, 2008; Whitford et al., 2001).

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Furthermore, it is also well known that rumen anaerobic fungi and methanogens form stable co-cultures due to the fungal species having high production levels of H2 (Orpin and Joblin, 1997). This leads to increased production of ATP and a metabolic shift towards enhanced acetate production in the rumen. Furthermore, the relationship enhances the production of fungal extracellular enzymes due to decreased catabolic repression caused by the methanogenic partner, which leads to enhanced rates of substrate breakdown in the rumen (Orpin and Joblin, 1997). Methanogenic archaea belonging to the genera Methanobacterium, Methanobrevibacter and Methanosphaera have also been found to be associated with H2-producing rumen ciliate protozoa either as endosymbionts or in commensal relationships with these protists (OheneAdjei et al., 2007). This is due to the ability of protozoal cells to produce hydrogenosomes, which the methanogens can then associate with at levels of 103–104 methanogens/protozoan (Ohene-Adjei et al., 2007; Tokura et al., 1997). It has also been established that these relationships are highly specialised and that only certain symbiotic rumen methanogens associate with ciliated protozoan in the Ophryoscolecidae family. However, the converse is true for free-living methanogens, and experimental data has recently highlighted that they are incapable of forming symbiotic relationships with rumen protists (Ohene-Adjei et al., 2007). Overall, given the rich microbial diversity in this ecosystem, it comes as no surprise that a number of microbial interactions should occur. However, while few, if any, acetogenic and SRB are known to exist in the rumen – it appears that competition, or more accurately co-existence, utilizing H2/CO2 as the main substrate under standard conditions is an activity undertaken mainly by methanogens in this ecosystem (Kamra, 2005; Liu and Whitman, 2008; Morvan et al., 1996).

4

Research Needs

While both the diversity and ubiquity of methanogens is surprising given their narrow metabolic characteristics, there are still significant gaps in our knowledge on these microbes in terms of the mammalian ecosystems.

4.1

Human Methanogenic Archaea and Disease

In the human colon and other ecosystems (mouth and vagina), studies have begun to intimate that methanogens are either useful biomarkers for diseases such as ulcerative colitis and Crohns disease, or have been associated with specific diseases such as vaginosis and periodontal and endodontic disease (Belay et al., 1990; Lepp et al., 2004; Scanlan et al., 2008; Vianna et al., 2006). Furthermore, a positive association between populations of Mbb. smithii and patients suffering from diverticulosis has been established (Weaver et al., 1986). However, it is not clear whether Koch’s postulates have been fulfilled to allow the etiology of any of these diseases to be ascribed to methanogenic archaea. In a number of cases, co-isolation/colocation of methanogenic archaea has occurred (Belay et al., 1990; Vianna et al., 2006). In other cases they are seen as promoters of disease such as periodontitis (Lepp et al., 2004).

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However to date, no virulence factors have been identified or associated with methanogenic archaea (Eckberg et al., 2003; Gophna et al., 2004). Given the lack of virulence factors identified from genomes of methanogenic archaea, and the general lack of fulfillment of Koch’s postulates – at this stage it is too early to state categorically whether methanogenic archaea have a significant role in human disease. Perhaps, further studies on the analysis of methanogenic archaeal genomes (especially of genes with currently unknown function) and isolation of more methanogenic archaeal phenotypes and any syntrophic microbial partners from sites of disease in mammalian gastrointestinal ecosystems will provide insight on this topic. One final area around methanogen associations with human disease is that of the relationship between the removal or lack of methanogens in the human colon and the putative association with a rare disorder – pnematosis cystoids coli (PCC). This disease is characterized by the presence of gas-filled cysts in the walls and mesentery of the large intestine (Florin, 1997). Research in the late 1980s and in the 1990s indicated that a significant correlation could be drawn in terms of the use of alkyl halides (chemically similar to chloroform) as a sedative in patients and the onset of PCC (Florin, 1997). Furthermore, both in vitro work using human feces and in vivo animal studies indicated that inhibition of methanogenesis and so-called ‘‘super H2 production’’ would lead to a counterperfusion supersaturation process occurring with H2 being transferred across the gut lumen and into the colon walls and mesenteric tissues (Florin, 1997). However, due to the growing trend of prescribing alternative sedatives, alkyl halides are less common now and as a consequence PCC has become a very rare disease.

4.2

Methanogenic Community Structure, Succession and Methane Mitigation

Clearly methanogenic diversity particularly in the rumen ecosystem will require more studies to determine the extent of phylogenetic diversity and to elucidate the currently large, diverse uncultured archaeal population (‘‘RCC group’’) associated with this ecosystem. Furthermore, and as noted by Janssen and Kirs (2008), while there are relatively few major phylogenetic groups, it is certainly not clear about the temporal effect on population stability or succession, and it is conceivable that in removing a major methanogenic group from the rumen ecosystem, a less abundant species may take over the ecological niche and predominate. More work is also required on not only identifying methanogenic species in human and rumen gastrointestinal ecosystems but also determining the factors affecting both succession and co-existence of major methanogenesis groups and species that are predominant in the rumen microbial ecosystem (See > Chapter 51, Vol. 1, Part 8). The utilization of modern molecular biology tools will clearly expedite and facilitate these questions being answered and aid in the production of robust rumen methane mitigation strategies.

5

Summary

This chapter has provided an overview of methanogenic communities associated with mammalian gastrointestinal systems. For detailed analyses and examination the reader is directed to

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some of the more recent articles and reviews (Janssen and Kirs, 2008; Liu and Whitman, 2008; Scanlan et al., 2008). As noted previously, while the ecosystems discussed here are not high lipid-content, senso stricto, they are part of larger biological systems that are either significantly impacted by lipid or are continuously exposed to lipids that are both simple and complex in nature. While the level of community diversity is significantly different between the ecosystems discussed i.e., to date, the major human colonic species Mbb. smithii has not been found to be associated with the rumen – the predominant methanogens in each gastrointestinal ecosystem are all clustered within the genus Methanobrevibacter. However, it is clear that further ecological studies will still be required to gain significant advances in our understanding of the methanogenic community structure and diversity in gastrointestinal ecosystems. Furthermore, more studies are needed to determine the effect of both host (diet, genetics, physiology) and environmental factors upon the methanogenic communities within mammalian gastrointestinal tract ecosystems.

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Mammalian Digestive Tract population: an association of methanogenic bacteria and diverticulosis. Gut 27: 698–704. Whitford MF, Teather RM, Forster RJ (2001) Phylogenetic analysis of methanogens from the bovine rumen. BMC Microbiol 1: 5. Wolin MJ, Miller TL, Stewart CS (1997) Microbemicrobe interactions. In The Rumen Microbial Ecosystem, 2nd edn. PN Hobson and CS Stewart (eds.). London: Chapman & Hall, pp. 467–491.

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56 Methanogenesis in the Digestive Tracts of Insects A. Brune Department of Biogeochemistry, Max Planck Institute for Terrestrial Microbiology, Marburg, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 708

2 2.1 2.2 2.3 2.4

Methane as a Product of Symbiotic Digestion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 709 Historical Aspects of Methane Emission by Insects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 709 Hydrogen is a Central Intermediate in Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 711 Association of Methanogens with Anaerobic Protists . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 713 Intercompartmental Transfer of Hydrogen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 714

3 3.1 3.2 3.3 3.4 3.5 3.6 3.7 3.8

Methanogenic Communities in Insect Guts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 715 Lower Termites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 715 Higher Termites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 715 Cockroaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 717 Scarab Beetles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 718 Non-Methanogenic Archaea . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 718 Methanogens Isolated from Insects . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 718 Relationship to Oxygen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 719 Absence of Methane Oxidation in Termites . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 721

4 Influence of Diet on Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 721 4.1 Do Methanogens Benefit the Insect? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 722 5

Insects as Source of Atmospheric Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 722

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_56, # Springer-Verlag Berlin Heidelberg, 2010

708

56

Methanogenesis in the Digestive Tracts of Insects

Abstract: Termites, cockroaches, and scarab beetles are the only insects known that emit methane, but they do so in impressive amounts. Methanogenesis occurs in the enlarged hindgut compartment and is fueled by hydrogen and reduced one-carbon compounds that are formed during the fermentative breakdown of plant fiber and humus. The methanogens typically colonize intestinal surfaces or occur as endosymbionts of protists. They fall into three major phylogenetic groups (Methanobacteriales, Methanosarcinales, and Methanomicrobiales) and form distinct phylogenetic clusters of mostly uncultivated archaea that are often unique to the intestinal tract of insects. The oxygen-reducing capacities of the few available isolates and their location in the microoxic gut periphery indicate that at least some species of this notoriously oxygen-sensitive group are well adapted to the continuous influx of oxygen into their habitat. Although colonization by methanogens seems to require a predisposition of the host, possible benefits for the insect remain to be discovered. The high methane emissions of termites, together with their enormous biomass especially in tropical regions, make them a significant natural source of this important greenhouse gas.

1

Introduction

Insects are the most successful class of terrestrial animals, with respect to both species richness and biomass. Most insects thriving on a fiber-rich diet have an enlarged hindgut that harbors a diverse microbiota of bacteria and protists, which participate in digestion (> Fig. 1). However,

. Figure 1 Structure of the intestial tract of a lower termite (a), a higher termite (b), and a scarab beetle larva (c). The crop (C) and the midgut (M) are free of methanogens. (a) In lower termites (here: Reticulitermes sp.), methanogens are located in a single, strongly dilated hindgut paunch (Pa) that tapers out via the colon into the rectum (R). (b) In most higher termites (here: a soil-feeding Cubitermes sp.), the hindgut is divided into several compartments (proctodeal segments P1–P5), all of which harbor methanogens. (c) In scarab beetle larvae (here: a humivorous Potosia sp.), the methanogens are exclusively found in the dilated hindgut (H) (Panel c reproduced from Werner, 1926a).

Methanogenesis in the Digestive Tracts of Insects

56

only certain groups of insects harbor methanogenic archaea. This chapter will review the diverse aspects of methanogenesis in insects, including the role of methanogens in symbiotic digestion and their location within the gut, the structure of the methanogenic community in the respective insect taxa, and the global relevance of methane emissions by insects. If this review appears biased towards termites, this simply reflects the limited number of studies on methanogenesis in other insects. The purpose of this chapter is to provide a broad overview of the area and to illustrate important concepts. For detailed information and an exhaustive survey of the literature, readers are referred to several excellent reviews focusing on different aspects of the subject (e.g., Breznak, 2000; Sugimoto et al., 2000; Brune, 2006; Hackstein et al., 2006; Purdy, 2007).

2

Methane as a Product of Symbiotic Digestion

2.1

Historical Aspects of Methane Emission by Insects

The first to suspect methane formation by termites was Sherburne F. Cook, an American physiologist, in the early 1930s. He had spent 2 years in Otto Warburg’s laboratory in Berlin and was interested in biological oxidations. While studying the respiratory gas exchange of the lower termite Zootermopsis nevadensis using Warburg manometry, he recognized that termites still formed substantial amounts of a gas other than CO2 when the oxygen in the vessel was depleted (Cook, 1932). He found that production of the unknown gas depended on the presence of the gut flagellates, which occupy the bulk of the hindgut in all phylogenetically lower termites (all families of Isoptera except Termitidae) and are essential for cellulose digestion. Cook was not able to analyze the gas, but he, inspired by the situation in ruminants, proposed that the gas was most likely hydrogen or methane, or a mixture of both. A few years later, Hungate (1938, 1939) confirmed the observations of Cook and documented hydrogen formation both for the gut flagellates of Zootermopsis and for living termites. Gilmour (1940b) noticed that a combustible gas was also formed (albeit in much smaller amounts) by the wood-feeding cockroach Cryptocercus punctulatus, which is a close relative of termites and which harbors the same type of gut flagellates. He added that the gas emitted by C. punctulatus did not consist exclusively of hydrogen because in almost every case some carbon dioxide was formed during combustion analysis in his modified Haldane apparatus. In the following years, Hungate (1943, 1946) conducted pioneering work on the fermentative metabolism of the gut flagellates of Zootermopsis species, which led to the first tenable concept of lignocellulose digestion in termite guts. Although he could show the conversion of cellulose to acetate, carbon dioxide, and hydrogen as major products, he never recognized that part of the hydrogen formed by the flagellates is subsequently metabolized to methane by other members of the gut microbiota (Hungate, 1977). Methane formation by termites was finally recognized more than 40 years after Cook’s initial observation of the unknown gas. The discovery was – as so often in science – a classical case of serendipity. During their studies of nitrogen fixation in termites (Breznak et al., 1973), Breznak and coworkers identified methane as an additional peak in the gas chromatographic assay of acetylene reduction to ethylene (personal communication), which they used to demonstrate nitrogenase activity in the living animals. The first publication on methanogenesis by insects was their following report on the closely related wood-feeding cockroach C. punctulatus, which also mentions methane production by termites (Breznak et al., 1974).

709

710

56

Methanogenesis in the Digestive Tracts of Insects

Their work led to the recognition that the amount of methane produced by these insects, most likely originating from the gut microbiota, is rather large and, based on body weight, in the same order of magnitude as that of ruminants (Breznak, 1975). It was soon recognized that other insects also emit methane (Bracke et al., 1978; Bayon and Etie´vant, 1980), but this capacity appeared to be restricted to relatively few groups of terrestrial arthropods: termites, cockroaches, scarab beetles, and millipedes (Hackstein and Stumm, 1994; > Table 1). Methane production was found among almost all termite species investigated, albeit at different levels (e.g., Brauman et al., 1992; Shinzato et al., 1992; Wheeler et al., 1996; Bignell et al., 1997; Sugimoto et al., 1998b). However, not all termite colonies were colonized by methanogens, and there was no specific trend in methane emission rates even among members of the same genus. In the closely related cockroaches, methane formation was apparently widespread, but not present in all lineages; also in millipedes, it was found only in certain taxa (Hackstein and Stumm, 1994). Almost all scarab beetles tested formed methane, but not all subfamilies were represented. The restriction of methanogenesis to certain phylogenetic groups of insects and the failed attempts to permanently establish methanogens in

. Table 1 Methane emission rates of invertebrates in comparison to that of cows and humans. Values are given in nmol g 1 h 1 and are based on fresh weight. S is the number of species tested (in the averages, only methane-emitting species were included). Data stem from studies using different experimental setups and were compiled from various sources (see footnotes)

Taxonomic group

Methane emission (range)

Diet

c

Methane emission (average)

S

Litter

0–415

58

6

Varied

0–268

46

9

Wood

0–1300

200

17

Macrotermitinaea,b,d,e

Wood, litter

20–670

170

12

Termitinae (wood-feeders)a,e

Wood, grass

40–210

120

11

Termitinae (soil-feeders)a,d,e

Soil, humus

150–1090

440

23

Nasutitermitinaea,b,d,e

Wood, grass

40–240

140

10

Apicotermitinaed

Soil, humus

50–700

280

7

0–741

255

7

580–960

780

Diplopoda Blattidaec Isoptera

Lower termitesa, b, e Higher termites

Scarabaeidae (Cetoniinae)

c

Cowsf

Humus Grass

Humansg a

Varied b

0–120 c

8 d

Data from Brauman et al. 1992; Shinzato et al. 1992; Hackstein and Stumm 1994; Bignell et al. 1995; eSugimoto et al. 1998; fKinsman et al. 1995 (assuming a body weight of 1400 kg); gBond et al. 1971 (calculated from pulmonary emissions, assuming a body weight of 75 kg and a colonic absorption of 20%)

Methanogenesis in the Digestive Tracts of Insects

56

non-methanogenic lineages by artificial infection suggests the presence of a genetic determinant and consequently a hereditary basis of the association (Hackstein et al., 1996, 2006).

2.2

Hydrogen is a Central Intermediate in Methanogenesis

Methane is formed almost exclusively by methanogenic archaea in two fundamentally different processes: (1) the reduction of CO2 or other C1 compounds to CH4 via the C1 pathway (hydrogenotrophic methanogenesis); and (2) the cleavage of acetate to CH4 and CO2 via the acetyl-CoA pathway (aceticlastic methanogenesis) (Hedderich and Whitman, 2006; Liu and Whitman, 2008). In most anoxic freshwater environments, both processes are present, resulting in the complete conversion of complex organic matter to CH4 and CO2. However, in intestinal environments, such as the rumen or the human gut, methane is mostly formed by hydrogenotrophic methanogens (Lange et al., 2005; Liu and Whitman, 2008) – there is no evidence for aceticlastic methanogenesis in insect guts. It is assumed that the relatively slowgrowing aceticlastic species cannot cope with the short retention times of intestinal habitats (Liu and Whitman, 2008), but it remains to be explained why they do not avoid washout by attaching to intestinal surfaces (see below). The most important electron donors of hydrogenotrophic methanogenesis are hydrogen and reduced C1 compounds (e.g., methanol and formate), which are formed during the

. Figure 2 Insects feeding on a fiber-rich diet typically harbor a dense microbiota of bacteria and often also protists in their enlarged hindgut. The products of symbiotic digestion (acetate and other shortchain fatty acids) are an important carbon and energy source for the host. In several insect groups, hydrogen formed by fermenting bacteria or protists is a major substrate for homoacetogenic bacteria and methanogenic archaea. The microorganisms colonizing the microoxic hindgut periphery have to cope with a constant influx of oxygen and are responsible for maintaining the anoxic status of the gut lumen (Brune, 2003, 2006).

711

712

56

Methanogenesis in the Digestive Tracts of Insects

fermentative breakdown of organic matter (> Fig. 2). Werner (1926a) noticed that copious amounts of hydrogen are formed during cellulose digestion in hindgut contents of scarab beetle larvae; he also isolated a strictly anaerobic bacterium catalyzing this reaction (Werner, 1926b). In lower termites, hydrogen is a major fermentation product of the cellulolytic gut flagellates (see above); it can accumulate to substantial concentrations (Ebert and Brune, 1997; Pester and Brune, 2007). Also phylogenetically higher termites (family Termitidae) accumulate and emit hydrogen (Sugimoto et al., 1998b; Schmitt-Wagner and Brune, 1999); since they lack gut flagellates, the gas is most likely formed by fermenting bacteria. Also many other insects emit substantial amounts of hydrogen (including non-methane-emitting taxa), and individuals from methane-emitting taxa that fail to produce methane often emit hydrogen instead (Hackstein and Stumm, 1994). There is abundant evidence for a pivotal role of hydrogen in methanogenesis in insect guts. Methane emission by lower termites strictly depends on the presence of (hydrogen-producing) gut flagellates (Odelson and Breznak, 1983; Rasmussen and Khalil, 1983; Messer and Lee, 1989). Methanogenesis in intact guts or gut homogenates of termites, cockroaches, and scarab beetle larvae is stimulated by the supply of external hydrogen (Brauman et al., 1992; Schmitt-Wagner and Brune, 1999; Lemke et al., 2001, 2003). When the cockroach Periplaneta americana is fed with metronidazole, a drug that inhibits hydrogen formation via pyruvateferredoxin oxidoreductase in both anaerobic bacteria and protists, methane emission is almost completely abolished (Bracke et al., 1978). When lower termites are fed with antibacterial drugs, both hydrogen and methane evolution rates increase strongly, which was the first indication that bacteria are competing with methanogens for the hydrogen formed by the flagellates (Odelson and Breznak, 1983). It turned out that CO2-reductive acetogenesis dominates over methanogenesis as the hydrogenotrophic process in most wood-feeding termites, whereas the opposite is true for most fungus-cultivating and soil-feeding termite species – both in gut homogenates and in situ (Breznak and Switzer, 1986; Brauman et al., 1992; Tholen and Brune, 1999, 2000; Pester and Brune, 2007). Despite the apparent competition for hydrogen, production and consumption of hydrogen are not tightly coupled, because termites (Zimmerman et al., 1982; Odelson and Breznak, 1983; Ebert and Brune, 1997; Sugimoto et al., 1998b; Schmitt-Wagner and Brune, 1999) and other methaneproducing insects (Hackstein and Stumm, 1994) also emit hydrogen in considerable amounts. At first sight, these phenomena are difficult to explain. For thermodynamic reasons, methanogens should outcompete homoacetogens for hydrogen, their common substrate (Cord-Ruwisch et al., 1988; Liu and Whitman, 2008), but this scenario is relevant only if the hydrogen partial pressures of the environment are around or below the H2 threshold concentrations for CO2-reductive acetogenesis by homoacetogens. However, microsensors studies revealed that hydrogen concentrations in termite guts are much higher than originally thought, indicating that sources and sinks of hydrogen are not evenly distributed within a gut (> Fig. 3a). Radial profiles showed high concentrations of hydrogen in the gut lumen and steep gradients towards the gut periphery in the hindgut of both lower and higher termites (Ebert and Brune, 1997; Schmitt-Wagner and Brune, 1999; Pester and Brune, 2007). In the wood-feeding lower termite Reticulitermes flavipes, the gut wall is densely colonized with methanogens (Leadbetter and Breznak, 1996; Leadbetter et al., 1998). This agrees with the observation that the strong hydrogen sink at the gut wall of this termite is caused by an anaerobic process (Ebert and Brune, 1997) and explains the apparent hydrogen limitation of methanogens in situ (Leadbetter and Breznak, 1996). The hydrogen concentration differences between the lumen and the periphery may also explain why methanogenesis is strongly stimulated

Methanogenesis in the Digestive Tracts of Insects

56

. Figure 3 Radial gut sections of a lower termite (a) and a cockroach (b), and axial gut section of a scarab beetle larva (c), illustrating the location of methanogens(), the radial concentration profiles of hydrogen and oxygen in agarose-embedded guts (in a, b), or the transfer of soluble reductants via the hemolymph (arrow in c). (a) In the hindgut (H) of lower termites, hydrogen production by gut flagellates leads to steep hydrogen gradients from the anoxic lumen towards the microoxic hindgut periphery. Methanogens are either associated with gut flagellates (fla) or attached to the hindgut wall, where they represent a major sink of hydrogen and possibly also of oxygen. (b) In cockroaches, methanogens are either associated with hydrogen-producing ciliates (cil) or attached to the hindgut cuticle, whose surface is enlarged by cuticular hairs. Methanogenesis is driven to a substantial part by a cross-epithelial transfer of hydrogen from the midgut (M) compartment. (c) In scarab beetle larvae, methanogens are attached to the gut epithelium or to tree-like epithelial invaginations (pseudosetae, ps). Hydrogen does not accumulate, but methanogenesis in the hindgut may be supported by formate (Fo) or other reductants produced in the midgut. For more details, see text.

by hydrogen supplied from the outside, whereas reductive acetogenesis – presumably located within the gut proper – is not (Tholen and Brune, 2000).

2.3

Association of Methanogens with Anaerobic Protists

In many anoxic environments, methanogens are associated with anaerobic protists (see van Hoek et al., 2000; Hackstein et al., 2001). Since many observations indicate that the methanogens benefit from an interspecies hydrogen transfer from the eukaryotic partner, sometimes even resulting in a mutual advantage (Schink, 1997), it is not astonishing that also the gut flagellates of termites are often associated with methanogens. Odelson and Breznak (1985) were first to note that a putatively axenic culture of a gut flagellate isolated from a Zootermopsis species contained a methanogenic symbiont. Lee et al. (1987), who exploited the characteristic blue-green fluorescence of cofactor F420 to localize the methanogens in the hindgut of Zootermopsis angusticollis, found that the methanogens are typically associated only with the smaller flagellates; the larger species, which are the major hydrogen source, appeared to lack methanogenic symbionts (Messer and Lee, 1989). In view of the high hydrogen concentrations throughout the gut lumen of almost all lower termites investigated (Pester and

713

714

56

Methanogenesis in the Digestive Tracts of Insects

Brune, 2007), it seems unlikely that methanogens are hydrogen-limited even if their particular host flagellate does not produce hydrogen itself (> Fig. 3a). Although colonization of certain gut flagellates by methanogens is not uncommon among lower termites (Shinzato et al., 1992; Hackstein and Stumm, 1994; Radek, 1994, 1997; Tokura et al., 2000; Hara et al., 2004), it is not a universal feature. In some termites, the methanogens are exclusively attached to the hindgut cuticle or to filamentous bacteria colonizing the gut wall (Hackstein and Stumm, 1994; Leadbetter and Breznak, 1996; Leadbetter et al., 1998); for others, a simultaneous colonization of gut wall and flagellates has been reported (Shinzato et al., 1992; Tokura et al., 2000). Since methanogens located in the gut periphery are clearly hydrogen-limited (see above), Sugimoto et al. (1998b) suggested that the rates of hydrogen and methane emission of different colonies may depend on the specific location of methanogens relative to the hydrogen source. The high methane emission rates of certain cockroaches may also be explained by a close association of methanogens with the hydrogen source. Most of the methane emitted by P. americana seems to be produced by the methanogens in Nyctotherus ovalis (Gijzen et al., 1991), an anaerobic ciliate present also in other cockroaches and certain millipedes (Hackstein and Stumm, 1994). Electron microscopy revealed that the endosymbiotic methanogens in the cytoplasm of the ciliate are located directly next to its hydrogenosomes (Gijzen et al., 1991; van Hoek et al., 2000), which would facilitate hydrogen transfer and is in agreement with the low hydrogen partial pressures and the severe hydrogen limitation of methanogenesis in the hindgut of cockroaches (see below). Also the small flagellates in the cockroach Supella supellectilium contain intracellular methanogens, whereas the flagellates present in many scarab beetles are devoid of such symbionts (Hackstein and Stumm, 1994).

2.4

Intercompartmental Transfer of Hydrogen

In lower termites, methanogens may be separated from the hydrogen sources in the gut lumen by their restriction to the hindgut periphery. In other termites, hydrogen production and methanogenesis may be located even in different compartments. In the hindgut of soil-feeding termites (Cubitermes spp.), hydrogen (presumably of bacterial origin) accumulates only in the anterior compartments, whereas methanogenic capacities are highest in the posterior, less alkaline gut regions (Schmitt-Wagner and Brune, 1999; Kappler and Brune, 2002). Since methanogenesis in these compartments depends strongly on external hydrogen, SchmittWagner and Brune (1999) postulated a cross-epithelial transfer of hydrogen between anterior and posterior gut segments, which are in close contact in vivo. Such cross-epithelial transfer of reducing equivalents seems to occur also in other methane-producing insects whose intestinal tracts show a functional compartmentalization. In the hindguts of Blaberus sp. and P. americana, which do not accumulate hydrogen, methanogenesis is severely hydrogen limited (Lemke et al., 2001; Sprenger et al., 2007). Methane production in ligated hindgut compartments comes close to the methane emission rates of living cockroaches only if external hydrogen is supplied or if the hindgut is incubated in the same vial as the hydrogen-producing midgut. When midgut and hindgut are placed in direct contact (mimicking the situation in vivo), steep hydrogen concentration gradients document a strong flux of hydrogen between the compartments (Lemke et al., 2001; > Fig. 3b). Methanogenesis in the posterior hindgut of soil-feeding termites (Cubitermes spp.) is not only stimulated by hydrogen but also by formate (Schmitt-Wagner and Brune, 1999),

Methanogenesis in the Digestive Tracts of Insects

56

and in the larval hindgut of the scarab beetle Pachnoda ephippiata, by formate and methanol (Lemke et al., 2003). The presence of considerable concentrations of formate in other gut compartments and in the hemolymph led to the suggestion that methanogenesis in the hindgut may be driven also by an intercompartmental transfer of reducing equivalents via the hemolymph (Lemke et al., 2003; > Fig. 3c).

3

Methanogenic Communities in Insect Guts

Information on the absolute numbers of methanogens in insect guts is scarce. Most-probablenumber determinations and viable counts indicate that R. flavipes harbors about 106 methanogens per gut, which would represent about 5% of the total prokaryote cell count (Leadbetter and Breznak, 1996; Tholen et al., 1997). These numbers are of limited value, however, since the methanogens colonize only the inner surface of the gut wall, and they may be also biased owing to the numerous bacterial endosymbionts of the gut flagellates, which occupy the bulk of the hindgut volume ( Table 2). Three Methanobrevibacter species have been isolated from the hindgut of R. flavipes (Leadbetter and Breznak, 1996; Leadbetter et al., 1998), and numerous other lineages of Methanobrevibacter-related sequences have been detected by cultivation-independent, 16S-rRNA-based surveys of numerous other termite species (Ohkuma et al., 1995; Ohkuma and Kudo, 1998; Ohkuma et al., 1999; Shinzato et al., 1999, 2001). These studies have documented the consistent presence of unique Methanobrevibacter-related phylotypes in each termite investigated, which comprise representative species of almost all families of lower termites. Many lower termites harbor more than one lineage of Methanobrevibacter, and selective cloning of archaeal 16S rRNA genes from capillary-picked suspensions of gut flagellates revealed that the phylotypes associated with the flagellates are phylogenetically distinct from those associated with the gut epithelium (Tokura et al., 2000). The Methanobrevibacter phylotypes associated with distantly related flagellates form a monophyletic cluster, which indicates that each of the Methanobrevibacter lineages within the same termite may have a preference for a particular microhabitat (Tokura et al., 2000; Hara et al., 2004; Inoue et al., 2008).

3.2

Higher Termites

The gut of higher termites is characterized by the absence of cellulolytic flagellates and shows (with the exception of the fungus-cultivating species) also a pronounced compartmentation,

715

(+)e

+

+

(+)d

Methanomicrobiales

(+)e

+

+

+

+

Thermoplasmatales

+

Halabacterials

Archaeal community structures of representatives of two subfamilies were analyzed

No data

Archaeal community structures of several species were analyzed (all soil feeders)

Archaeal community structure of one species was analyzed

One species studied, very few clones analyzed

Almost all families of lower termites are represented

Archaeal community structures of two wood-feeding species was analyzed

Only the endosymbionts of Nyctotherus spp. were systematically studied

Only the endosymbionts of Nyctotherus spp. were systematically studied

Comments

a

All clones represent Methanobrevibacter species, bMany cockroach families harboring methane-producing taxa remain to be investigated, cSo far only represented by the isolate Methanomicrococcus blatticola (see > Table 3), dOnly a single clone was detected (Shinzato et al., 1999), eOnly detected in the humivorous Pachnoda ephippiata (Cetoniinae), but not in the phytophagous Melolontha melolontha (Melolonthinae)

Scarab beetles

+

+

Termitinae

Apicotermitinae

+

+

Nasutitermitinae

+

+

+

(+)c

Macrotermitinae

Higher termites

Lower termites

+

(+)

Blattellidae

Termites

+

+

Blattidae

Cockroachesb

Millipedes

Methanosarcinales

56

Methanobacterialsa

. Table 2 Major phylogenetic lineages of Euryarchaeota detected in the hindgut of insects and millipedes, mostly by 16S-rRNA-based analysis. Parentheses indicate that sequences were not detected in all taxa. In some cases, the apparent absence of a lineage may be due to poor diversity coverage of the analysis (see also comments). The information was compiled from numerous references; for details, see text

716 Methanogenesis in the Digestive Tracts of Insects

Methanogenesis in the Digestive Tracts of Insects

56

which goes hand in hand with a remarkable dynamics of intestinal pH and redox potential (Brune, 2006). The methanogenic community is much more diverse than that of lower termites. Not a single strain has been isolated in pure culture, but 16S-rRNA-based analyses revealed the presence of Methanomicrobiales, Methanosarcinales, and Methanobacteriales in almost all species investigated, representing three of the four subfamilies of higher termites (family Termitidae; > Table 2). This holds true for the wood-feeding Nasutitermes takasagoensis (Nasutitermitinae) (Ohkuma et al., 1999; Miyata et al., 2007) as well as the soil-feeding species Pericapritermes nitobei (Ohkuma et al., 1999), Cubitermes orthognathus (Friedrich et al., 2001), and Cubitermes fungifaber (Donovan et al., 2004) (all Termitinae). The fungus-cultivating Odontotermes formosanus (Macrotermitinae) may be an exception because only Methanosarcinales were recovered (Ohkuma et al., 1999). Although undersampling cannot be excluded, since only a limited number of clones were investigated, the result is supported by the study of Brauman et al. (2001), who reported that Methanosarcinales may account for the total archaeal community in the fungus-growing Macrotermes subhyalinus (Macrotermitinae). Representatives of the fourth subfamily (Apicotermitinae), which are mostly soil-feeding, have not been investigated. The clones of Methanomicrobiales and Methanosarcinales recovered from representatives of the different subfamilies of higher termites are phylogenetically closely related and form characteristic clusters with the phylotypes obtained from other insects. The clones of Methanobacteriales all fall into the phylogenetic radiation of the genus Methanobrevibacter, but seem to represent several distinct lineages, just as their relatives in the lower termites (see Dighe et al., 2004; Miyata et al., 2007; Purdy, 2007). A detailed analysis of the archaeal community structure in the different gut compartments of C. orthognathus showed that the different phylogenetic groups are not evenly distributed among the different compartments (Friedrich et al., 2001). Methanosarcinales colonize the anterior, extremely alkaline hindgut compartment (P1), whereas Methanobacteriaceae and Methanomicrobiales predominate in the posterior compartments (P3/4a and P4b). The diversity of methanogens obtained with clone libraries is in general agreement with the results obtained by the rRNA-based dot-blot analysis (Brauman et al., 2001). Using groupspecific probes, they detected Methanobacteriales in the guts of almost all termite species studied, regardless of its diet or taxonomic classification, whereas Methanosarcinales were detected in about half of the species. However, there are also obvious inconsistencies between their dataset and the results obtained with clone libraries (e.g., the failure to obtain any signal for Methanomicrobiales), which indicate that the results should be interpreted with caution. Nevertheless, the large discrepancies between the combined hybridization signals of the groupspecific probes and the archaeal domain probe obtained with almost all termites investigated may also reflect the presence of other, non-methanogenic archaea (see below).

3.3

Cockroaches

Archaeal clones obtained from the hindgut of the wood-feeding cockroaches Panesthia angustipennis and Salganea taiwanensis (Blaberidae; Hara et al., 2002) represent the same phylogenetic lineages as those obtained from wood-feeding termites, but the community structure differs considerably. While the archaeal communities in termites are dominated either by Methanobacteriales or by Methanomicrobiales, the majority of clones obtained from cockroaches are Methanosarcinales. Nevertheless, they are closely related to the

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Methanosarcinales clones obtained from all higher termites and to Methanomicrococcus blatticola, an isolate from P. americana (Sprenger et al., 2000), which corroborates that also this group of methanogens is widely distributed in insects. Interestingly, the endosymbiotic methanogens of Nyctotherus spp., anaerobic ciliates common in the hindgut of many cockroaches and certain millipedes, are not related to the endosymbiotic Methanomicrobiales colonizing other heterotrichous ciliates living in sediments. They form yet another lineage in the genus Methanobrevibacter (van Hoek et al., 2000), distinct from the Methanobrevibacter lineages present in termites or mammalian guts (Dighe et al., 2004). Apparently, the different ciliate lineages have independently acquired free-living methanogens as endosymbionts during adaptation to their respective ecological niches (van Hoek et al., 2000).

3.4

Scarab Beetles

The archaeal community in the hindgut of scarab beetle larvae is dominated by distinct lineages of Methanobrevibacter-related clones (Egert et al., 2003, 2005) that are closely related to the Methanobrevibacter species in termite guts. While no other methanogens are present in the phytophagous Melolontha melolontha larva, the humivorous P. ephippiata larva harbors also a smaller fraction of Methanosarcinales (> Table 2), which are again closely related to M. blatticola and the clones obtained from termites and cockroaches (see above).

3.5

Non-Methanogenic Archaea

A substantial fraction of archaeal clones obtained from the hindguts of humivorous beetle larvae (Egert et al., 2003), soil-feeding and wood-feeding termites (Friedrich et al., 2001; Miyata et al., 2007), and wood-feeding cockroaches (Hara et al., 2002) form a deeply branching clade among the Thermoplasmatales (> Table 2). The phylotypes obtained from the different insects represent closely related yet distinct lineages and cluster with a few other clones of intestinal origin (rumen, swine manure). Miyata et al. (2007) observed that the proportion of Thermoplasmatales-related clones in N. takasagoensis increased substantially when the termite was fed a diet consisting only of xylan. However, before any inferences can be made regarding the so-far unknown metabolic properties of this novel lineage, it remains to be determined whether this relative shift was caused by an absolute increase of Thermoplasmatales or a decrease of the methanogenic populations. Donovan et al. (2004) detected several clones affiliated with Halobacteriales in the hindgut of the soil-feeding C. fungifaber. Archaeal clone libraries of higher termites (Friedrich et al., 2001; Miyata et al., 2007) and scarab beetle larvae (Egert et al., 2003) contained also a number of clones affiliated with Crenarchaeota.

3.6

Methanogens Isolated from Insects

Very few methanogens from insect guts have been obtained in pure culture (> Table 3). Three of the four isolates are from the lower termite R. flavipes, where they colonize the hindgut cuticle or are attached to filamentous bacteria associated with the gut wall (Leadbetter

Methanogenesis in the Digestive Tracts of Insects

56

and Breznak, 1996; Leadbetter et al., 1998). As other Methanobrevibacter species isolated from the human gut or the rumen, the isolates show a very restricted substrate spectrum, growing almost exclusively on H2 + CO2. The fourth isolate is M. blatticola, so far the only methanogenic archaeon isolated from a cockroach and the first cultivated representative of a novel lineage of Methanosarcinales associated with insect guts. It is specialized in the obligately hydrogen-dependent reduction of methanol or methylamines to methane (Sprenger et al., 2000). Its strict requirement for hydrogen in methanogenesis is explained by its inability to oxidize methyl groups to carbon dioxide (Sprenger et al., 2005). The substrate affinities of M. blatticola for hydrogen and methanol are higher than those of other methylotrophic methanogens (Methanosphaera stadtmanae, Methanosarcina barkeri), and since the use of methanol as the terminal electron acceptor is thermodynamically more favorable than the use of carbon dioxide, M. blatticola may have a competitive advantage over other methanogens at low hydrogen concentrations (Sprenger et al., 2007).

3.7

Relationship to Oxygen

Since methanogenesis is an oxygen-sensitive process, it is not astonishing that it is restricted to the hindgut, the only compartment of the insect gut exhibiting a negative redox potential (Bayon, 1980; Ebert and Brune, 1997; Kappler and Brune, 2002). Like all other methanogens, the four isolates mentioned above are obligate anaerobes, i.e., pure cultures do not grow in media containing even traces of oxygen (Liu and Whitman, 2008). Notably, both M. blatticola and the three Methanobrevibacter species isolated from termites specifically colonize the hindgut epithelium of their host (Leadbetter and Breznak, 1996; Leadbetter et al., 1998; Sprenger et al., 2000). Here, the organisms are at the bottom end of the radial hydrogen gradient of the gut proper, but may benefit from external hydrogen entering the hindgut by cross-epithelial transfer from other compartments (Schmitt-Wagner and Brune, 1999; Lemke et al., 2001; Sprenger et al., 2007). In view of the proverbial oxygen sensitivity of methanogens, their adherence to the hindgut epithelium – a location potentially exposing them to inflowing oxygen (Ebert and Brune, 1997; Brune and Friedrich, 2000) – is particularly intriguing. However, recent biochemical and genomic studies have revealed the capacity of methanogens to reduce molecular oxygen. The oxygen-reducing activity of Methanobrevibacter arboriphilus is mediated by a F420H2 oxidase (FprA; Seedorf et al., 2004); homologues of the fprA gene are present also in the termite gut isolate M. cuticularis (A. Brune, unpublished results). This may explain why the termite gut isolates, as well as the closely related Methanobrevibacter arboriphilus, are able to remove inflowing oxygen from agar gradient tubes (Leadbetter and Breznak 1996; Tholen et al., 2007) and to maintain an anoxic status and thus the metabolic activity of dense cell suspensions exposed to controlled oxygen fluxes (Tholen et al., 2007). Like other Methanobrevibacter species, the termite gut isolates possess both catalase and superoxide dismutase (Leadbetter and Breznak, 1996; Leadbetter et al., 1998), which would protect the enzymes of hydrogenotrophic methanogenesis, particularly the extremely oxygen-sensitive H2-forming methylenetetrahydromethanopterin dehydrogenase (Hmd), from inactivation by reactive oxygen species produced endogenously or by neighboring microbes in situ (Seedorf et al., 2004). Shima et al. (2001), stimulated by the results of Leadbetter and Breznak (1996), showed the presence of a heme-containing catalase in Methanobrevibacter arboriphilus, but only when cells were grown in media containing hemin. If catalase is protective for the Methanobrevibacter spp.

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. Table 3 Methanogenic archaea isolated from insect guts Order/Species

Source

Substrates

Reference

Methanobacteriales Methanobrevibacter cuticularis

Reticulitermes flavipes

H2 + CO2a

Leadbetter and Breznak 1996

Methanobrevibacter curvatus

Reticulitermes flavipes

H2 + CO2

Leadbetter and Breznak 1996

Methanobrevibacter filiformis

Reticulitermes flavipes

H2 + CO2

Leadbetter et al., 1998

Periplaneta americana

H2 + CH3OH b Sprenger et al., 2000

Methanosarcinales Methanomicrococcus blatticola a

Slow growth also on formate Growth also on methylamines in the presence of H2; no growth on H2 + CO2

b

in situ, its synthesis will depend upon acquisition of a suitable precursor of heme biosynthesis from the hindgut fluid, stemming from the degradation of heme proteins in the gut and/or from other microorganisms in the hindgut community. It is not clear how much oxygen the Methanobrevibacter species colonizing the gut periphery of termites experience in situ. Colonization of the hindgut wall by methanogens is patchy (Leadbetter and Breznak, 1996), and it cannot be excluded that the methanogens are restricted to less-oxygenated microsites. Interestingly, there is a considerable discrepancy between the amount of methane emitted by R. flavipes and the potential capacity of its methanogenic population (Leadbetter and Breznak, 1996). Since a mitigation of methane emissions by aerobic methane-oxidizing bacteria can be ruled out (see below), the low rates of methanogenesis in the hindgut might indicate that the methanogens themselves redirect the flow of reducing equivalents from methanogenesis towards oxygen reduction, in order to maintain an oxygen status that allows survival and growth (Tholen et al., 2007). The survival of methanogens in the hindgut periphery may be facilitated by the regulation of tracheal gas exchange, by which insects limit the tracheal oxygen partial pressure in order to guard tissues against oxidative damage (Lighton, 1996). It has been shown that Z. nevadensis can restrict the spiracular area to limit oxygen influx via the trachea, which should also affect the oxygen status of the gut microenvironment (Lighton and Ottesen, 2005). A discontinuous gas exchange, which has been reported for cockroaches and scarab beetle larvae (Bijnen et al., 1996), could lead to temporal phases of anoxia in the gut periphery of these insects. Nevertheless, the location of methanogens at the gut wall of lower termites remains enigmatic. It has been suggested that an attachment to the hindgut cuticle may protect from predation or prevent washout from the gut, which may compensate methanogens for the negative effects of hydrogen limitation and exposure to inflowing oxygen (Breznak, 2000). In that case, an association with gut flagellates that maintain a stable position in the anoxic and hydrogen-rich hindgut lumen should be of considerable advantage, not only for methanogens but also for obligately anaerobic bacteria. In many higher termites, cockroaches, scarab beetle larvae, and millipedes, a major fraction of the prokaryotic community, including many

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F420-fluorescent cells, adheres to cuticular structures projecting from the hindgut wall into the lumen (Bignell et al., 1980; Hackstein et al., 2006). Such cuticular differentiations are not specific for insects harboring methanogens; similar structures are also found in crickets and locusts. Ultrastructural studies indicate that they may play other roles in symbiotic digestion, e.g., in the resorption of microbial fermentation products by the host (see Hackstein et al., 2006 for a detailed and excellently illustrated treatise on this subject).

3.8

Absence of Methane Oxidation in Termites

The constant influx of oxygen and the presence of methane should render insect guts an ideal habitat for aerobic methane-oxidizing bacteria. However, there is no report documenting the presence or activity of methanotrophic bacteria within the intestinal tract of insects. Moreover, several pieces of evidence corroborate the absence of methane oxidation from termites. Methane emissions of Zootermopsis (Messer and Lee, 1989) and several other termite species from different families (Pester et al., 2007) are not, or only slightly, affected by the absence of oxygen, and all attempts to amplify the pmoA gene, a marker for particulate methane monooxygenase, from hindgut DNA extracts of these termites were unsuccessful (Pester et al., 2007). Moreover, 14CH4 added to the headspace of vials containing living termites (R. flavipes or C. orthognathus) was not converted to 14CO2 (Pester et al., 2007). This eliminates the gut itself as a potential sink for this important greenhouse gas, which leaves methane oxidation in mound material and surrounding soil as a potentially important factor mitigating methane production by termites at the environmental level (see below).

4

Influence of Diet on Methanogenesis

The distribution pattern of methane-emitting insect taxa in the tree of life suggests that colonization by methanogens requires certain prerequisites in the form of a fiber-rich diet and a negative redox potential of the hindgut (Hackstein et al., 1996). However, there are only a few studies of dietary influences on methane production by insects. In the cockroach P. americana, methane emission varies under different feeding regimens (Gijzen et al., 1991; Zurek and Keddie, 1998). Differences in the pattern of fermentation products indicated that methanogenesis is the major electron sink in the hindgut of individuals fed on high-fiber diet, whereas a low-fiber diet favors reductive acetogenesis and the accumulation of formate (Kane and Breznak, 1991; Zurek and Keddie, 1998). The situation in termites is different. Here, methane emission rates are less pronounced in wood-feeding and fungus-cultivating species, but increase from wood–soil interface feeders to true soil feeders (Wandiga and Mugedo, 1987; Brauman et al., 1992; Shinzato et al., 1992; Rouland et al., 1993; Bignell et al., 1997; Sugimoto et al., 1998b) – a feeding guild that thrives exclusively on peptide-rich soil organic matter (Ji and Brune, 2006). It is tempting to suggest that these differences are diet related, but information on fermentative processes in the hindguts of humivorous insects is sparse, and also the biology of their rather diverse methanogenic community (see below) has to be better understood before a reasonable hypothesis can be proposed; the availability of hydrogen alone is clearly not the answer.

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Methanogenesis in the Digestive Tracts of Insects

Do Methanogens Benefit the Insect?

There is no evidence that colonization by methanogens increases the fitness of an insect, but the number of studies addressing this question is limited. Although all lower termites die of starvation after removal of their cellulolytic gut flagellates, elimination of methanogens from all flagellates by feeding of Z. angustacollis with bromoethanesulfonic acid (BES) did not affect the survival of the termites (Messer and Lee, 1989). Individuals of the cockroach P. americana reared in the absence of protozoa grew more slowly, had a lower adult body weight, and did not produce methane (Gijzen and Barugahare, 1992). Suppression of methanogenesis by addition of BES to the drinking water of cockroaches containing protozoa shifted the fermentation pattern towards an increased propionate formation, but had no effects on insect body weight or the number of N. ovalis cells (Gijzen and Barugahare, 1992). Zurek and Keddie (1998) confirmed that BES treatment greatly reduced methane production but had no significant effect on weight gain and development time of P. americana, but also found that VFA concentrations in the hindgut were not significantly altered.

5

Insects as Source of Atmospheric Methane

Owing to the possible role of ‘‘greenhouse gases’’ in radiative forcing of the atmosphere, early publications on methane formation by termites (see above) soon aroused the interest of atmospheric chemists. Zimmerman et al. (1982) recognized that the considerable methane emission rates of these animals, multiplied by their huge numbers especially in tropical ecosystems, result in a potential source of considerable strength. Their first estimate of global methane emissions by termites (150 Tg year 1) was enormous and was immediately criticized by other workers for the non-representative choice of termite species and an overestimation of termite biomass and food consumption (Rasmussen and Khalil, 1983; Collins and Wood, 1984). In the following years, numerous studies followed up on this subject, increasing the accuracy of the available dataset and decreasing the estimates of global methane emissions by termites by at least an order of magnitude (for critical reviews, see Sanderson, 1996; Bignell et al., 1997; Sugimoto et al., 1998a). Nevertheless, even current estimates of methane production by termites are still far from accurate. There are enormous differences in the methane emission of termite taxa, whether they are based on the individual, body mass, or their ratio to carbon dioxide production. However, less than 3% of all termite species have been investigated, and the contribution of individual taxa to termite biomass in different ecological regions is often not clear (Sanderson, 1996). Since methane emission by termites is strongly correlated to the mode of nutrition (see above), global estimates now incorporate scaling factors to account at least for the ratio of soil feeders to non-soil feeders in a given region (Bignell et al., 1997). Predictions are further confounded by the strong intercolony variation of methane emissions and by large monthly and diurnal fluctuations caused by temperature and humidity changes (e.g., Wheeler et al., 1996; Sugimoto et al., 1998b), which are usually not accounted for in the existing datasets. Most measurements on methane emission by termites have been conducted in glass jars containing a number of termites taken from the nest. Among all biases introduced by this setup, the most problematic one is the failure to account for methane oxidation within the mound material or the surrounding soil. Field measurements indicated that the net emissions of methane from termite nests into the atmosphere are often much smaller than methane

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. Figure 4 The relative contributions of termites and other sources to the global atmospheric budget of methane, distinguishing natural sources and those under anthropogenic influence. Data from Kvenvolden and Rogers (2005), based on the third assessment report of the IPCC (Ehhalt et al., 2001).

production by the termite population in the nest (e.g., Seiler et al., 1984; Khalil et al., 1990; Delmas et al., 1992; Bignell et al., 1997; MacDonald et al., 1998, 1999; Eggleton et al., 1999). However, the importance of this process varies. Up to 40% of the methane produced within mounds of Coptotermes lacteus is oxidized before it reaches the atmosphere (Khalil et al., 1990), whereas the mound material of Macrotermes jeanneli shows no significant methane oxidation capacity (Darlington et al., 1997), underlining the need to introduce emission factors into the global estimates that account for the effect of methane oxidation in different taxa and/or feeding guilds. Since many termites build either huge or often diffuse nests, it is extremely tedious or even impossible to determine the population of termites in a colony and its net emission of methane into the atmosphere. It was therefore an important conceptual advance when Sugimoto et al. (1998a) estimated the proportion of methane oxidized en route to the atmosphere by comparing the carbon isotope ratios (13C/12C) of methane produced by the termites and of methane emitted from the mounds. Owing to the preferential conversion of lighter methane molecules by methane-oxidizing bacteria, the residual methane becomes enriched in heavier carbon, and the extent of this shift, combined with the fractionation factor of methane oxidation, allows calculation of the emission factor of the respective colony. The results of this study were striking because they convincingly documented that – contrary to the prediction from laboratory data – net emissions of methane from the colonies of soil-feeding termites are much lower than those of wood-feeding termites. Based on a comprehensive dataset of termite biomass and methane emission rates compiled from their own studies and from the literature, and taking into account the emission factors determined for the different feeding guilds, Sugimoto et al. (2000) concluded that the contribution of termites to global methane emissions is probably less than 10 Tg year 1 (1.5–7.4 Tg; Sugimoto et al., 1998b) but almost certainly below 20 Tg year 1 – the number that is still used in the last global budget presented by the IPCC (Denman et al., 2007).

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Termites may still represent the second largest natural source of methane on the planet, but their contribution to the total source strength (ca. 600 Tg year 1) is certainly dwarfed by the sources under anthropogenic influence (> Fig. 4). The contribution of methane emission by soil invertebrates other than termites has received only little attention. Based on their laboratory measurements and on rough estimates for the biomass distribution of higher arthropod taxa in tropics and subtropics, Hackstein and Stumm (1994) pointed out that millipedes and scarab beetles may represent another globally important source of methane. Also here, further studies are clearly needed.

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Hungate RE (1943) Quantitative analyses of the cellulose fermentation by termite protozoa. Ann Entomol Soc Am 36: 730–739. Hungate RE (1946) The symbiotic utilization of cellulose. J Elisha Mitchell Sci Soc 62: 9–24. Hungate RE (1977) The rumen microbial ecosystem. Annu Rev Microbiol 33: 1–20. Inoue J, Noda S, Hongoh Y, Ui S, Ohkuma M (2008) Identification of endosymbiotic methanogen and ectosymbiotic spirochetes of gut protists of the termite Coptotermes formosanus. Microb Environ 23: 94–97. Ji R, Brune A (2006) Nitrogen mineralization, ammonia accumulation, and emission of gaseous NH3 by soilfeeding termites. Biogeochemistry 78: 267–283. Kane MD, Breznak JA (1991) Effect of host diet on production of organic acids and methane by cockroach gut bacteria. Appl Environ Microbiol 57: 2628–2634. Kappler A, Brune A (2002) Dynamics of redox potential and changes in redox state of iron and humic acids during gut passage in soil-feeding termites (Cubitermes spp.). Soil Biol Biochem 34: 221–227. Khalil MAK, Rasmussen RA, French JRJ, Holt JA (1990) The influence of termites on atmospheric trace gases: CH4, CO2, CHCl3, N2O, CO, H2, and light hydrocarbons. J Geophys Res 95: 3619–3634. Kvenvolden KA, Rogers BW (2005) Gaia’s breath – global methane exhalations. Marine Petroleum Geol 22: 579–590. Lange M, Westermann P, Kiær Ahring B (2005) Archaea in protozoa and metazoa. Appl Microbiol Biotechnol 66: 465–474. Leadbetter JR, Breznak JA (1996) Physiological ecology of Methanobrevibacter cuticularis sp. nov. and Methanobrevibacter curvatus sp. nov., isolated from the hindgut of the termite Reticulitermes flavipes. Appl Environ Microbiol 62: 3620–3631. Leadbetter JR, Crosby LD, Breznak JA (1998) Methanobrevibacter filiformis sp. nov., a filamentous methanogen from termite hindguts. Arch Microbiol 169: 287–292. Lee MJ, Schreurs PJ, Messer AC, Zinder SH (1987) Association of methanogenic bacteria with flagellated protozoa from a termite hindgut. Curr Microbiol 15: 337–341. Lemke T, van Alen T, Hackstein JHP, Brune A (2001) Cross-epithelial hydrogen transfer from the midgut compartment drives methanogenesis in the hindgut of cockroaches. Appl Environ Microbiol 67: 4657–4661. Lemke T, Stingl U, Egert M, Friedrich MW, Brune A (2003) Physicochemical conditions and microbial activities in the highly alkaline gut of the humusfeeding larva of Pachnoda ephippiata (Coleoptera:

Scarabaeidae). Appl Environ Microbiol 69: 6650–6658. Lighton JRB (1996) Discontinuous gas exchange in insects. Annu Rev Entomol 41: 309–324. Lighton JRB, Ottesen EA (2005) To DGC or not to DGC: oxygen guarding in the termite Zootermopsis nevadensis (Isoptera: Termopsidae). J Exp Biol 208: 4671–4678. Liu Y, Whitman WB (2008) Metabolic, phylogenetic, and ecological diversity of the methanogenic archaea. Ann N Y Acad Sci 1125: 171–189. MacDonald JA, Eggleton P, Bignell DE, Forzi F, Fowler D (1998) Methane emission by termites and oxidation by soils, across a forest disturbance gradient in the Mbalmayo Forest Reserve, Cameroon. Global Change Biol 4: 409–418. MacDonald JA, Jeeva D, Eggleton P, Davies R, Bignell DE, Fowler D, Lawton J, Maryati M (1999) The effect of termite biomass and anthropogenic disturbance on the CH4 budgets of tropical forests in Cameroon and Borneo. Global Change Biol 5: 869–879. Messer AC, Lee MJ (1989) Effect of chemical treatments on methane emission by the hindgut microbiota in the termite Zootermopsis angusticollis. Microb Ecol 18: 275–284. Miyata R, Noda N, Tamaki H, Kinjyo K, Aoyagi H, Uchiyama H, Tanaka H (2007) Phylogenetic relationship of symbiotic archaea in the gut of the higher termite Nasutitermes takasagoensis fed with various carbon sources. Microb Environ 22: 157–164. Odelson DA, Breznak JA (1983) Volatile fatty acid production by the hindgut microbiota of xylophagous termites. Appl Environ Microbiol 45: 1602–1613. Odelson DA, Breznak JA (1985) Nutrition and growth characteristics of Trichomitopsis termopsidis, a cellulolytic protozoan from termites. Appl Environ Microbiol 49: 614–621. Ohkuma M, Kudo T (1998) Phylogenetic analysis of the symbiotic intestinal microflora of the termite Cryptotermes domesticus. FEMS Microbiol Lett 164: 389–395. Ohkuma M, Noda S, Horikoshi K, Kudo T (1995) Phylogeny of symbiotic methanogens in the gut of the termite Reticulitermes speratus. FEMS Microbiol Lett 134: 45–50. Ohkuma M, Noda S, Kudo T (1999) Phylogenetic relationships of symbiotic methanogens in diverse termites. FEMS Microbiol Lett 171: 147–153. Pester M, Brune A (2007) Hydrogen is the central free intermediate during lignocellulose degradation by termite gut symbionts. ISME J 1: 551–565. Pester M, Tholen A, Friedrich MW, Brune A (2007) Methane oxidation in termite hindguts: absence of

Methanogenesis in the Digestive Tracts of Insects evidence and evidence of absence. Appl Environ Microbiol 73: 2024–2028. Purdy KJ (2007) The distribution and diversity of euryarchaeota in termite guts. Adv Appl Microbiol 62: 63–80. Radek R (1994) Monocercomonides termitis n. sp., an oxymonad from the lower termite Kalotermes sinaicus. Arch Protistenkunde 144: 373–382. Radek R (1997) Spirotrichonympha minor n. sp., a new hypermastigote termite flagellate. Eur J Protistol 33: 361–374. Rasmussen RA, Khalil MAK (1983) Global production of methane by termites. Nature 301: 700–702. Rouland C, Brauman A, Labat M, Lepage M (1993) Nutritional factors affecting methane emission from termites. Chemosphere 26: 617–622. Sanderson MG (1996) Biomass of termites and their emissions of methane and carbon dioxide: A global database. Global Biogeochem Cycles 10: 543–557. Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61: 262–280. Schmitt-Wagner D, Brune A (1999) Hydrogen profiles and localization of methanogenic activities in the highly compartmentalized hindgut of soil-feeding higher termites (Cubitermes spp.). Appl Environ Microbiol 65: 4490–4496. Seedorf H, Dreisbach A, Hedderich R, Shima S, Thauer RK (2004) F420H2 oxidase (FprA) from Methanobrevibacter arboriphilus, a coenzyme F420-dependent enzyme involved in O2 detoxification. Arch Microbiol 182: 126–137. Seiler W, Conrad R, Scharffe D (1984) Field studies of methane emission from termite nests into the atmosphere and measurements of methane uptake by tropical soils. J Atmos Chem 1: 171–186. Shima S, Sordel-Klippert M, Brioukhanov A, Netrusov A, Linder D, Thauer RK (2001) Characterization of a heme-dependent catalase from Methanobrevibacter arboriphilus. Appl Environ Microbiol 67: 3041–3045. Shinzato N, Yoshino H, Yara K (1992) Methane production by microbial symbionts in the lower and higher termites of the Ryukyu Archipelago. In Endocytobiology V. S Sato, M Ishida, H Ishikawa (eds.). Tu¨bingen: Tu¨bingen University Press, pp. 161–166. Shinzato N, Matsumoto T, Yamaoka I, Oshima T, Yamagishi A (1999) Phylogenetic diversity of symbiotic methanogens living in the hindgut of the lower termite Reticulitermes speratus analyzed by PCR and in situ hybridization. Appl Environ Microbiol 65: 837–840. Shinzato N, Matsumoto T, Yamaoka I, Oshima T, Yamagishi A (2001) Methanogenic symbionts and

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the locality of their host lower termites. Microb Environ 16: 43–47. Sprenger WW, van Belzen MC, Rosenberg J, Hackstein JHP, Keltjens JT (2000) Methanomicrococcus blatticola gen. nov., sp., nov., a methanol- and methylamine-reducing methanogen from the hindgut of the cockroach Periplaneta americana. Int J Syst Evol Microbiol 50: 1989–1999. Sprenger WW, Hackstein JHP, Keltjens JT (2005) The energy metabolism of Methanomicrococcus blatticola: physiological and biochemical aspects. Antonie v Leeuwenhoek 87: 289–299. Sprenger WW, Hackstein JH, Keltjens JT (2007) The competitive success of Methanomicrococcus blatticola, a dominant methylotrophic methanogen in the cockroach hindgut, is supported by high substrate affinities and favorable thermodynamics. FEMS Microbiol Ecol 60: 266–275. Sugimoto A, Inoue T, Kirtibutr N, Abe T (1998a) Methane oxidation by termite mounds estimated by the carbon isotopic composition of methane. Global Biogeochem Cycles 12: 595–605. Sugimoto A, Inoue T, Tayasu I, Miller L, Takeichi S, Abe T (1998b) Methane and hydrogen production in a termite-symbiont system. Ecol Res 13: 241–257. Sugimoto A, Bignell DE, MacDonald JA (2000) Global impact of termites on the carbon cycle and atmospheric trace gases. In Termites: Evolution, Sociality, Symbioses, Ecology. T Abe, DE Bignell, M Higashi (eds.). Dordrecht: Kluwer, pp. 409–435. Tholen A, Brune A (1999) Localization and in situ activities of homoacetogenic bacteria in the highly compartmentalized hindgut of soil-feeding higher termites (Cubitermes spp.). Appl Environ Microbiol 65: 4497–4505. Tholen A, Brune A (2000) Impact of oxygen on metabolic fluxes and in situ rates of reductive acetogenesis in the hindgut of the wood-feeding termite Reticulitermes flavipes. Environ Microbiol 2: 436–449. Tholen A, Schink B, Brune A (1997) The gut microflora of Reticulitermes flavipes, its relation to oxygen, and evidence for oxygen-dependent acetogenesis by the most abundant Enterococcus sp. FEMS Microbiol Ecol 24: 137–149. Tholen A, Pester M, Brune A (2007) Simultaneous methanogenesis and oxygen reduction by Methanobrevibacter cuticularis at low oxygen fluxes. FEMS Microbiol Ecol 62: 303–312. Tokura M, Ohkuma M, Kudo T (2000) Molecular phylogeny of methanogens associated with flagellated protists in the gut and with the gut epithelium of termites. FEMS Microbiol Ecol 33: 233–240. van Hoek AHAM, van Alen TA, Sprakel VSI, Leunissen JAM, Brigge T, Vogels GD, Hackstein JHP (2000) Multiple acquisition of methanogenic archaeal

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symbionts by anaerobic ciliates. Mol Biol Evol 17: 251–258. Wandiga SO, Mugedo JAZ (1987) Methane emissions by tropical termites feeding on soil, wood, grass, and fungus combs. Kenya J Sci 8A: 19–25. Werner E (1926a) Die Erna¨hrung der Larve von Potosia ¨ kol cuprea Fbr. (Cetonia floricola Hbst.). Z Morph O Tiere 6: 150–206. Werner E (1926b) Der Erreger der Zelluloseverdauung bei der Rosenka¨ferlarve (Potosia cuprea Fbr.) Bacillus cellulosam fermentans n. sp. Zbl Bakter II 67: 297–330.

Wheeler GS, Tokoro M, Scheffrahn RH, Su N-Y (1996) Comparative respiration and methane production rates in nearctic termites. J Insect Physiol 42: 799–806. Zimmerman PR, Greenberg JP, Wandiga SO, Crutzen PJ (1982) Termites: A potentially large source of atmospheric methane, carbon dioxide, and molecular hydrogen. Science 218: 563–565. Zurek L, Keddie BA (1998) Significance of methanogenic symbionts for development of the American cockroach, Periplaneta americana. J Insect Physiol 44: 645–651.

Part 1

Introduction: Theoretical Considerations

1 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization F. Widdel . F. Musat Max Planck Institute for Marine Microbiology, Bremen, Germany [email protected] [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 732

2

Some Basic Thermodynamic Aspects of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . 732

3 Energetics of Hydrocarbon Utilization by Microorganisms . . . . . . . . . . . . . . . . . . . . . . . 738 3.1 Catabolic Net Reactions of Hydrocarbons from the Energetic Perspective . . . . . . . . . 738 3.2 Hydrocarbon Activation from the Energetic Perspective . . . . . . . . . . . . . . . . . . . . . . . . . . . . 743 4 Quantitative Aspects of Cell Synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747 4.1 ATP and Growth Yields . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 747 4.2 Requirement for Minerals (N, P, Fe) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 751 5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 752 Appendix . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 753

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_57, # Springer-Verlag Berlin Heidelberg, 2010

732

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Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

Abstract: Hydrocarbons represent ‘‘energy-rich’’ growth substrates for aerobic microorganisms and in principle allow high growth yields. In contrast, the energy gain with hydrocarbons in many anaerobic microorganisms is very low. The maximum gain of energy per mol of hydrocarbon degraded in the catabolism is predicted from calculated DG values. Some anaerobic degradation reactions of hydrocarbons with very low energy gain as well as anaerobic activation reactions of hydrocarbons deserve particular attention from a bioenergetic point of view.

1

Introduction

The study of microbial growth with hydrocarbons and their degradation often gets into energetic aspects, even though at a glance the metabolism of hydrocarbons is not basically different from that of other organic compounds. The overall metabolism in a chemotrophic organism follows the universal bifurcate carbon flow: One part of the carbon substrate together with sources of other elements (N, P, S, Fe, etc.) is used for synthesis of cell components, a process referred to as anabolism (synthetic metabolism, assimilation). The anabolic ‘‘upgrading’’ of the substrate requires and dissipates much energy, which is usually provided in the form of ATP and derived from another part of the carbon substrate. This part of the substrate necessarily undergoes degradation; the degradative substrate flow is referred to as catabolism (energy metabolism, dissimilation). Still, there are some energetic peculiarities in the metabolism of hydrocarbons which deserve attention. (1) First, even though flammability of hydrocarbons at the air implies ‘‘energy richness,’’ they are not energy-rich under all circumstances. In the absence of oxygen, hydrocarbons are less energy-rich than for instance the less flammable glucose. Whereas the latter provides energy for various modes of fermentative growth, fermentation of saturated, aromatic and many other unsaturated non-aromatic hydrocarbons is energetically not feasible1; this is one reason why they tend to be preserved in deep reservoirs. (2) Second, hydrocarbons are chemically unreactive at room temperature. Their use in the metabolism has to begin with an activation reaction, the introduction of a functional group, which may require and ‘‘waste’’ energy from the overall energy budget of the microorganism. Also energies of transition states in the activation reactions have been of interest for a mechanistic understanding. (3) Third, for the theoretical treatment of energy conservation with hydrocarbons as well as for the estimation of microbial cell mass involved in hydrocarbon (petroleum) bioremediation, growth yields (cell mass produced per amount of hydrocarbon utilized) are of interest. This chapter briefly addresses some of these energetic peculiarities and quantitative aspects of hydrocarbon metabolism (> Fig. 1).

2

Some Basic Thermodynamic Aspects of Hydrocarbons

Hydrocarbons, the main constituents of oil and gas, are the major source of energy in our industrialized society. A prominent property of hydrocarbons is thus their ‘‘energy richness.’’ More precisely, this term expresses that energy is released if they are oxidized with oxygen, and that the amount of energy released per unit mass (the gravimetric energy density) of a liquid or solid hydrocarbon is higher than that from the oxidation of many other chemical

1

A fermentable hydrocarbon is, for instance, the unsaturated acetylene. Also some other unsaturated hydrocarbons are, at least theoretically, fermentable.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1

. Figure 1 The metabolism of hydrocarbons in chemotrophic microorganisms follows the universal bifurcate substrate flow into cell synthesis and degradation. A peculiarity in comparison to the metabolism of most non-hydrocarbon substrates is the activation which may require and dissipate energy.

compounds or elements (Appendix > Table 5). In the case of gaseous hydrocarbons, a high volumetric energy density is obvious if compared to that of other gases (Appendix > Table 5). This ‘‘energy richness’’ is due to the high affinity of the two constituents, hydrogen and carbon, for oxygen, and to the absence of oxidized carbon groups (such as C
OH or C=O groups). The low atomic masses of hydrogen and carbon2 is another factor that contributes to the high gravimetric energy density. It is the high gravimetric energy density which, together with the abundance of hydrocarbons in the form of petroleum, has made them ideal fuels for vehicles and aircrafts. Another technical advantage is the formation of volatile products (CO2, H2O vapor). Feasibility and maximum energy gains of formulated stoichiometric reactions are expressed by their free energy changes, the G-values. If a reaction is feasible under the given conditions (exergonic reaction), the DG-value is negative by convention. A positive value necessarily indicates that the reaction can in principle not occur under the given conditions (endergonic reaction), and a value of zero indicates that reactants and products are in equilibrium. Most reactions in chemistry and biology are associated with liberation of heat 2

H, 1.008; C, 12.011.

733

734

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Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

to the surroundings (exothermic reactions), which is expressed by their heat or enthalpy3 changes (DH-values). Some reactions consume heat from the surroundings (endothermic reactions), and a few of such type also occur in microorganisms. Free energy or enthalpy changes are calculated from free energies of formation (DGf , sometimes also termed Gf ) or enthalpies of formation (DHf , sometimes also termed Hf ), respectively, which are given for standard conditions4 and for which there is a broad data basis. Appendix > Table 6 compiles the values under standard conditions for several hydrocarbons and a number of other compounds which often appear in catabolic reactions. For a reaction a Aþb B!c Cþd D

ð1Þ

(with a, b, c, d being the stoichiometric factors), the standard free energy change (viz. for all compounds at standard conditions) is the difference

 ð2Þ DG  ¼ c DGf C þ d DGf D
a DGf A þ b DGf B Calculation of the free energy change DG for non-standard activities (a, in case of gases termed fugacity; a must not be confused with the stoichiometric factor a) considers the ‘‘nonchemical’’ energy change associated with dilution or concentration (‘‘volume work’’) of each component. These are logarithmic functions involving the gas constant and absolute temperature, the sum of which modifies the free energy change for standard activities, DGStandard, according to DG ¼ DG Standard þ R T ln

aCc  aDd aAa  aBb

ð3Þ

T in this equation must be the temperature for which the underlying DGStandard value has been given (viz. usually 298.15 K), and DG values at other temperatures cannot be calculated by this equation.5 The activities (effective concentrations) of solutes, a, can be usually substituted with acceptable precision by the actual concentrations in mol l
1; similarly, the fugacities (effective pressures) of gases can be substituted by their pressures in atm, an otherwise obsolete unit.6 With such simplification, as well as with R = 8.315  10
3 kJ K
1 mol
1, 3

Heat change of reaction under constant pressure. T = 298.15 K (25  C); standard activity of solutes, a = 1; standard (partial) pressure of gases = 101 kPa (standard fugacity = 1). 5 DGStandard values at temperatures other than 298.15 K can be calculated via the integrated ‘‘Delta-version’’ of the Gibbs-Helmholtz equation   @ DG
DH ¼ : @T T p T2 4

Assuming that temperature dependence of DH within the range of physiologically relevant temperatures is negligible, the free energy change at temperatures other than 298.15 K (but at standard activities) is   T T DGTStandard ¼ 1
: DG  þ DH  298:15 298:15 The same result is obtained from DG ¼ DH
T DS by assuming that DH and DS are essentially constant within the range of physiologically relevant temperatures. 6 The apparent correctness of the old unit atm is due to the fact that it is numerically equivalent with standard fugacity = 1. Activities and fugacities are by definition without units, and the formally correct approximated ½AActual substitution would be aA ¼ , etc. Here, use of the modern unit Pa or kPa for [A] etc. is coherent. ½AStandard

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1

T = 298.15 K (25  C), the common use of kJ as energy unit, and ln x = 2.303 lg x, (3) converts to DG ¼ DG  þ 5:71 lg

½Cc ½Dd ½Aa ½Bb

ðat 298:15 KÞ

ð4Þ

Hydrocarbons in the aqueous surroundings of microorganisms can be often considered with good approximation to have the activities of their gaseous, liquid or solid standard states, viz. aHydrocarbon = 1, or [Hydrocarbon] = 1. For instance, if a gaseous hydrocarbon at standard pressure dissolves in water and reaches the dissolution equilibrium (DG of transfer = 0), it is thermodynamically treated like the gas, even though the dissolved concentration is in the range of 10
3 M (Appendix > Fig. 6). The same holds true for liquid hydrocarbons: Despite the extremely low saturation concentration of long-chain alkanes in water, the hydrocarbon dissolved in water has the activity (strictly speaking the chemical potential) of the pure liquid hydrocarbon phase. The free energy data (Appendix > Table 6) reveal some basic and sometimes ‘‘counterintuitive’’ thermodynamic properties of hydrocarbons. Many hydrocarbons are metastable (thermodynamically unstable; DGf positive) with respect to the elements, even though decay into the elements is usually ‘‘kinetically inhibited.’’ In the case of acetylene (ethyne), however, compression at room temperature can trigger the release of the energy in a violent decay into the elements. For this reason, compressed welding acetylene in steel bottles must be stabilized by adsorption to a carrier such as acetone. But also hydrocarbons that are stable with respect to the elements (even the rather stable ethane) are metastable with respect to decay into native carbon and methane, the most stable hydrocarbon: 2 C2 H6 ! CGraphite þ 3 CH4 DG  ¼
43:3 kJ ðmol C2 H6 Þ
1

ð5Þ

In the presence of CO2 or bicarbonate, even methane is metastable: CH4 þ CO2 ! 2 CGraphite þ 2 H2 O DG  ¼
29:2 kJ ðmol CH4 Þ
1 þ CH4 þ HCO
3 þ H ! 2 CGraphite þ 3 H2 O

DG 0 ¼
33:9 kJ ðmol CH4 Þ
1

ð6Þ

ð7Þ

Nevertheless, formation of elemental carbon by reactions (5)–(7) is kinetically strongly inhibited and has not been observed in abiotic or biotic systems at room temperature. But once the element has been formed by geothermal metamorphism of buried biomass or petroleum (Tissot and Welte, 1984), it is the thermodynamically stable species of carbon as long as additional reducing or oxidizing components are absent. In the presence of a mild oxidant, not the element but rather CO2 and its ionic forms, HCO3
and CO32
, are the thermodynamically stable forms of carbon. Other forms or intermediate oxidation states (H2C=O, CO, HCOOH, C2-compounds, etc.), like all natural organic compounds, are metastable7 with respect to a conversion to CH4, CO2 and H2O, even without involvement of

7

The extremely low hypothetical equilibrium concentrations of these species can be calculated.

735

736

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

an oxidant or reductant. If on the other hand a reductant with negative enough redox potential is present, the only stable form of carbon is CH4. Again, intermediate oxidation states (CH3OH, reduced C2-compounds, etc.) are metastable. The stability ‘‘regions’’ of the mentioned carbon species are elegantly illustrated in a plot of the redox potential versus the pH (E-pH-diagram, Pourbaix diagram; > Fig. 2). Another thermodynamically interesting principle is revealed in the homologous series of n-alkanes. n-Alkanes become increasingly unstable with increasing chain length, whereas the

. Figure 2 Stability diagram (Pourbaix diagram) of carbon species. The equilibrium (borderline) redox potential E (in V) as a function of pH was calculated from DGf values (in J) according to P o P o Q DGf; red DGf; ox
0:0592 aox þ E ¼ lg Q nF ared n with n = number of electrons; F = 96,485 C mol
1; a = activity. Paox includes the H+-activity, the negative logarithm of which is the pH. Activities or fugacities: CO2, CH4, 1.0; HCO3
and CO32
, 10
2 (black) or 1.0 (gray). The borderlines between CO2 (g), HCO3
and CO32
in their standard states represent the pKa values. Note that the pKa1 value for CO2 (g) is 7.8 (vertical gray line), whereas that of CO2 (aq) is 6.35 (not shown here), the more commonly known one. The system H2O/H2 (electrochemically the same as 2H+/H2) is indicated for comparison.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1

heat of formation shows an opposite trend (> Fig. 3).8 The heat is the energy released during C
H-bond formation from the elements (even though such alkane formation is not observed in reality). Hence, the thermodynamically feasible disintegration of a long-chain alkane into its elements would consume heat: C16 H34 ! 16 CGraphite þ 17 H2 DG  ¼
49:8 kJ ðmol C16 H34 Þ
1 

DH ¼ þ454:4 kJ ðmol C16 H34 Þ

ð8Þ


1

This thermodynamically ‘‘allowed’’ cooling of the surroundings (and the system), which is a decrease in the entropy of the surroundings, is explained by the numerically higher entropy increase of the reacting system; the molecules of the gaseous H2 that are formed in high number carry a high amount of ‘‘hidden heat.’’9 Furthermore, the homologous n-alkane series

. Figure 3 Free energy and enthalpy (heat) of formation of alkanes from C1 to C18. The free energy of formation of C16 includes a literature value and the value extrapolated in this graph.

Linearity in the series of the higher alkanes may be a ‘‘pre-assumption’’ and basis for calculation of DGf or DHf values of compounds in homologous series via incremental additions. In the numerous sources of thermodynamic data, the original basis underlying such data is often difficult to trace back. 9 Also, the highly ordered (‘‘improbable’’) structure of the long-chain alkane contributes to thermodynamic instability. 8

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738

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Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

reveals the transition from gaseous to liquid hydrocarbons (n-butane/n-pentane), which is mirrored by a discontinuity of the DHf values. This is because liquid pentane has ‘‘given off ’’ the heat of condensation to the surroundings (liquid n-pentane, the real standard state: DHf =
173 kJ mol
1; hypothetical gaseous standard state: DHf =
146 kJ mol
1). The discontinuity of DGf is less pronounced. Liquid pentane as a highly volatile compound (boiling point, 36.2  C) is almost in equilibrium with the gaseous state (liquid n-pentane: DGf =
9.21 kJ mol
1; hypothetical gaseous standard state: DGf =
8.11 kJ mol
1).

3

Energetics of Hydrocarbon Utilization by Microorganisms

The biological utilization of a hydrocarbon can be examined bioenergetically (1) on the level of the net reaction performed by a microorganism and (2) on the level of individual enzymatic reactions. Among the latter, those of hydrocarbon activation are usually of highest interest and therefore briefly addressed in this overview.

3.1

Catabolic Net Reactions of Hydrocarbons from the Energetic Perspective

The G of a reaction (the ‘‘system’’) is the maximum amount of energy that a second system can theoretically conserve via coupling to this reaction under full reversibility. However, coupling can only proceed outside of the equilibrium, viz. if the overall reaction of the two systems is more or less irreversible and dissipates free energy. The actual amount of useful energy provided by the catabolic reactions is therefore always less than the calculated DG. The subsequent anabolism with many highly irreversible reactions then dissipates most of the free energy. > Table 1 lists generalized equations for the degradation of hydrocarbons, and > Table 2 several particular reactions with naturally important electron acceptors and the associated energy changes. The aerobic oxidation with O2 as electron acceptor provides biochemically the highest amount of energy, methanogenic degradation the lowest. Reactions with NO2
or N2O are even more exergonic than those with O2 (> Table 2 includes methane oxidation with nitrite as an investigated example; Ettwig et al., 2008). However, there is no evidence that the higher energy available with NO2
or N2O in comparison to O2 as electron acceptor is conserved; biochemically, O2 allows conservation of even more energy from the same amount of organic substrate. The theoretically higher energy gain is due to the ‘‘extra energy content’’ of NO2
and N2O with respect to O2: 4 NO2
+ 4 H+ ! 2 N2 + 3 O2 + 2 H2O, DG 0 =
116 kJ (mol NO2
)
1; 2 N2O ! 2 N2 + O2, DG  =
104 kJ (mol N2O)
1. One of the least exergonic catabolic reactions is the anaerobic oxidation of methane (> Table 2). Under certain environmental conditions, the net free energy change under in situ concentrations of the reactants may be only around DG =
20 kJ mol (CH4)
1 (Nauhaus et al., 2002). The fact that this minute amount is further shared between two organisms with different metabolism challenges the energetic understanding of energy conservation under ‘‘low-energy’’ conditions (viz. life at low chemical potential), a topic developed in the

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1

. Table 1 Generalized equations for the catabolism (dissimilation, degradation) of hydrocarbons with various electron acceptors, and for the anabolism (assimilation) of hydrocarbons into cell mass Catabolism 4 CcHh + (4c + h) O2 ! 4c CO2 + 2h H2O 10 CcHh + (8c + 2h) NO3
+ (8c + 2h) H+! 10c CO2 + (4c + h) N2 + (4c + 6h) H2O CcHh + (4c + h) Fe(OH)3 + (3c + h) CO2 ! (4c + h) FeCO3 + (6c + 2h) H2Oa 8 CcHh + (4c + h) SO42
+ (8c + 2h) H+ ! 8c CO2 + (4c + h) H2S + 4h H2O 8 CcHh + (8c
2h) H2O ! (4c
h) CO2 + (4c + h) CH4 Anabolism 4 CcHh + h CO2 + (4c
h) H2O ! (4c + h) hCH2Oi dass = 0.133/(4c + h) mol g
1 Yass = (4c + h)/0.133 g mol
1 17 CcHh + (4h
c) CO2 + (14c
5h) H2O ! (4c + h) C4H7O3 dass = 0.165/(4c + h) mol g
1 Yass = (4c + h)/0.165 g mol
1 17 CcHh + (4h
c) CO2 + (4c + h) NH3 + (10c
6h) H2O ! (4c + h) C4H8O2N dass = 0.166/(4c + h) mol g
1 Yass = (4c + h)/0.166 g mol
1 a

Because many reactions take place in environments containing inorganic carbon, reactions with Fe(OH)3 are for convenience written with the relatively insoluble FeCO3 (siderite) as a product

study of other syntrophic associations (Jackson and McInerney, 2002; Schink, 1997, 2002; chapter by McInerney et al. in this volume). Another anaerobic reaction of a hydrocarbon of thermodynamic interest is the conversion of alkanes to methane (Anderson and Lovley, 2000; Jones et al., 2008; Zengler et al., 1999), an endothermic reaction (for explanation see remark on (8)): 4 C16 H34 þ 30 H2 O ! 49 CH4 þ 15 CO2 ðgÞ DG  ¼
372 kJ ðmol C16 H34 Þ
1 

ð9Þ


1

DH ¼ þ206 kJ ðmol C16 H34 Þ

The Gibbs–Helmholtz equation predicts that the reaction becomes increasingly exergonic with increasing temperature (Dolfing et al., 2008). The process involves three organisms, (1) the hexadecane-degrading syntrophs (C16H34 + 16 H2O ! 8 CH3COO
+ 8 H+ + 17 H2), (2) acetate-cleaving microorganisms which are either methanogens (CH3COO
+ H+ ! CH4 + CO2) or additional syntrophs (CH3COO
+ H+ + 2 H2O ! 2 CO2 + 4 H2), and (3) H2-utilizing methanogens (CO2 + 4 H2 ! CH4 + 2 H2O). The available energy per transferred acetate, the ‘‘metabolic unit’’ in this syntrophism, is only around
47 kJ mol
1; this amount is shared between three organisms. The thermodynamic constraints of this reaction with respect to petroleum hydrocarbon conversion to methane have been examined (Dolfing et al., 2008).

739





928.3
344d

!5 CO2 (g) + 4 N2 (g) + 14 H2O

!3 CO2 (g) + 4 N2 (g) + 10 H2O

!8 FeCO3 + 14 H2O

3 CH4 (g) + 8 NO2
+ 8 H+

CH4 (g) + 8 Fe(OH)3 + 7 CO2 (g)

4 C2H6 (g) + 2 H2O

4 C2H6 (g) + 7

SO42


2 C2H6 (g) + 7 O2 (g)

Ethane

+ 14 H


73
36

!7 CH4 (g) + CO2 (g)


1,467

!8 CO2 (g) + 7 H2S (aq) + 12 H2O

!4 CO2 (g) + 6 H2O


16.6d,e

!HCO3
+ HS
+ H2O

CH4 (g) + SO42


+


21.2


16.5d,e

d

!HCO3
+ H2S (aq) + H2O

+2H

+

!CO2 (g) + H2S (aq) + 2 H2O

CH4 (g) + SO4

2


CH4 (g) + SO42
+ H+

5 CH4 (g) + 8 NO3 + 8 H

+


766.2


813.3d

+ H2O

CH4 (g) + 2 O2 (g)

+H


818.0d

!HCO3
+

!CO2 (g) + 2 H2O

Productsa

CH4 (g) + 2 O2 (g)

Methane

Reactantsa

Free energy change of reaction per mol hydrocarbonb, DG or DG0 (kJ mol
1)


2


38


1,560


11.3


33.5


20.9


559


993


788


903.0


890.4

115

117


310

18


57

1


723


215


73


301


243

Enthalpy change Entropy change of reaction of reaction per mol hydrocarbon, per mol hydrocarbonb,c, DH (kJ mol
1) DS or DS0 (J K
1 mol
1)

1

. Table 2 Thermodynamic characteristics of observed and some hypothetical reactions of selected hydrocarbons. The degradation of hydrocarbons to methane and carbon dioxide is often endothermic

740 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

2 C3H8 (g) + 5 SO4
63


372


3,008
1,423
214
135

!12 CO2 (g) + 6 H2O

!30 CO2 (g) + 15 N2 (g) + 30 H2O

!30 FeCO3 + 48 H2O

!24 CO2 (g) + 15 H2S (aq) + 12 H2O

!15 CH4 (g) + 9 CO2 (g)

2 C6H6 (l) + 15 O2 (g)

5 C6H6 (l) + 30 NO3
+ 30 H+

C6H6 (l) + 30 Fe(OH)3 + 24 CO2 (g)

4 C6H6 (l) + 15 SO42
+ 30 H+

4 C6H6 (l) + 18 H2O

Benzene
3,202


632

!64 CO2 (g) + 49 H2S (aq) + 68 H2O

+ 98 H

!49 CH4 (g) + 15 CO2 (g)

4 C16H34 (l) + 30 H2O

4 C16H34 (l) + 49 SO4

+

!80 CO2 (g) + 49 N2 (g) + 134 H2O

2



9,757

!32 CO2 (g) + 34 H2O

2 C16H34 (l) + 49 O2 (g)

5 C16H34 (l) + 98 NO3
+ 98 H+


10,392


137

!19 CH4 (g) + 5 CO2 (g)

4 C6H14 (l) + 10 H2O

n-Hexadecane


238

!12 CO2 (g) + 14 H2O

!24 CO2 (g) + 19 H2S (aq) + 28 H2O


4,023


117

!5 CH4 (g) + CO2 (g)


2,108

!6 CO2 (g) + 5 H2S (aq) + 8 H2O

!3 CO2 (g) + 4 H2O

4 C6H14 (l) + 19 SO42
+ 38 H+

+ 10 H

+

2 C6H14 (l) + 19 O2 (g)

n-Hexane

2 C3H8 (g) + 2 H2O

2


C3H8 (g) + 5 O2 (g)

Propane

71


7


2,026


2,883


3,268

206


50


9,445


10,701

66


33


4,163

6


46


2,220

691

696


2,024

419


220

1,937

1,952

1,047


1,036

682

688


471

232

236


375 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1 741

!9 CH4 (g) + 5 CO2 (g)

2 C7H8 (l) + 10 H2O


313
186

!10 CO2 (g) + 6 H2S (aq) + 4 H2O

!6 CH4 (g) + 4 CO2 (g)

C10H8 (c) + 8 H2O


2,247


4,782

C10H8 (c) + 6 SO42
+ 12 H+

C10H8 (c) + 48 Fe(OH)3 + 38 CO2 (g) !48 FeCO3 + 76 H2O

!10 CO2 (g) + 4 H2O

!50 CO2 (g) + 24 N2 (g) + 44 H2O

C10H8 (c) + 12 O2 (g)

5 C10H8 (c) + 48 NO3
+ 48 H+

186

61


3,171


4,541


5,156

96

2


2,421


3,449


3,910

1,245

1,253


3,098

808


212

801

806


2,456

473


293

Enthalpy change Entropy change of reaction of reaction per mol hydrocarbon, per mol hydrocarbonb,c, DH (kJ mol
1) DS or DS0 (J K
1 mol
1)

Indicated standard states: g, gaseous; l, liquid; aq, aqueous, dissolved in water; c, crystalline b If protons are involved, DG0 (viz. for [H+] = 10
7 M, pH = 7) is given DH
DG c Here calculated via DS ¼ T d Standard free energy changes of reactions formulated with CO2 differ from those formulated with HCO3
because the reaction HCO3
+ H + ! CO2 (g) + H2O is exergonic under standard conditions at pH = 7, with DG0 =
4.7 kJ mol
1 e Reactions with H2S and HS
as products are energetically equivalent because both sulfide species are essentially in equilibrium under standard conditions

a


142

!14 CO2 (g) + 9 H2S (aq) + 8 H2O

2 C7H8 (l) + 9 SO42
+ 18 H+


5,093


238

!36 FeCO3 + 58 H2O

C7H8 (l) + 36 Fe(OH)3 + 29 CO2 (g)

Naphthalene


1,689

!35 CO2 (g) + 18 N2 (g) + 38 H2O

5 C7H8 (l) + 36 NO3
+ 36 H+


3,823
3,590

!7 CO2 (g) + 4 H2O

C7H8 (l) + 9 O2 (g)

Toluene

Productsa

Free energy change of reaction per mol hydrocarbonb, DG or DG0 (kJ mol
1)

1

Reactantsa

. Table 2 (Continued)

742 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

3.2

1

Hydrocarbon Activation from the Energetic Perspective

As any chemical or biochemical reaction, the activation reaction of hydrocarbons involves two energetic aspects. These are the net G of the reaction (and its share in the overall catabolic DG), and the energy level which during the activation reaction is attained by the energy-rich short-lived transition state in the active site of the hydrocarbon-activating enzyme; an apparent transition state may further resolve into elementary reactions upon closer examination (> Fig. 4). Net free energy changes of several activation reactions of hydrocarbons are listed in > Table 3. The activation of a hydrocarbon by introduction of a functional group to allow further metabolic processing is usually not a problem from a merely thermodynamic point of view. For instance, an O2-independent hydroxylation of methane by dehydrogenation at a hypothetical ‘‘methane dehydrogenase’’ employing a mildly oxidizing biological agent such as cytochrome c (Cyt cox/Cyt cred, E  = E 0 = +0.245 V) would be thermodynamically allowed:

. Figure 4 Free energy changes during a fictive reaction of a hydrocarbon (S; in principle any other substrate) converted to an end product (P), free energy change of the activation reaction, and free energies of transition states at the activating enzyme. The scheme is a simplification because it does not display any electron acceptor that allows oxidation and the indicated free energy changes.

743

C6H6 + O2 ! [Intermediate; NADH-recycling] ! o-C6H4(OH)2

CH4 + CoM
S
S
CoB ! CoM
S
CH3 + HS
CoB

Benzene

g

Benzene

Carboxylationf

Butenee





4
31

C6H6 + Carrier
COO
! C6H5
COO
+ Carrier
H


7


31 to
35


35 to
39


27 to
31

+30


335

C6H6 + HCO3
! C6H5COO
+ HOH

CH3
CH2
CH=CH2 + H2O ! CH3
CH2
HCH
CH2OH

Addition of water to isolated double bond

C6H5CH3 +
OOC
CH=CH
COO
!
OOC
[C6H5CH2]CH
HCH
COO





R
CH2
CH3 + OOC
CH=CH
COO ! OOC
[(R)
CH
(CH3)]CH
HCH
COO




Toluene




CH4 +
OOC
CH=CH
COO
!
OOC
[CH3]CH
HCH
COO


n-Alkane

Methanee

Addition to fumarate

Methane

! C15H31
CH2OH + H2O + NAD


368

+

C15H31
CH3 + O2 + NADH + H +

Widdel et al. (2007)

Widdel et al. (2007)

Widdel et al. (2007)

Rabus et al. (2001)

Rabus et al. (2001)

Rabus et al. (2001)

Shima and Thauer (2005)

Widdel et al. (2007)

This article

Widdel et al. (2007)

Reference

b

Fate of one reactant is visualized in bold DG0 is indicated if protons are involved c Would be less exergonic with an electron donor of less negative redox potential than that of NAD+/NADH (
0.320 V) d Sum of the following formal reactions: C15H31
CH3 + 0.5 O2 ! C15H31
CH2OH, DG =
148.5 kJ mol
1; 0.5 O2 + NADH + H + ! H2O + NAD +, DG0 =
219.6 kJ mol
1 (calculated from DE0 = 1.138 V; Thauer et al., 1977) e Hypothetical reaction f Carboxylations have been suggested on the basis of chemical analyses g A carboxyl carrier and donor has not been suggested or identified. The present free energy change is based on a calculation with oxaloacetate as a purely hypothetical carboxyldonor that may be energetically comparable to potent carboxyl-donors such as carboxy-biotin

a

d


344c

Reactiona

CH4 + O2 + NADH + H + ! CH3OH + H2O + NAD+

Methyl-coenzyme M reductase

Benzene

n-Hexadecane

Methane

Mono- and dioxygenation

Type of activation; Compound

1

Free energy changeb DG or DG0 (kJ mol
1)

. Table 3 Free energies of activation reactions of saturated, monounsaturated and aromatic hydrocarbons. Also purely hypothetical reactions have been included. Values are given for standard conditions (pH = 7, if protons are involved)

744 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

    CH4 þ 2 Cyt c Fe3þ þ H2 O ! CH3 OH þ 2 Cyt c Fe2þ þ 2 Hþ DG 0 ¼
16 kJ ðmol CH4 Þ
1

1 ð10Þ

The problem lies in the high energy barrier, mainly due to the apolar and very stable C
H-bond that must be attenuated by an appropriate biocatalyst. Despite the astounding capabilities of enzymes to decrease energy barriers of chemically difficult reactions, there is not always the ideal biochemical solution to any activation problem. Not every thermodynamically possible but kinetically inhibited reaction can be catalyzed to take place at any rate.10 To overcome the activation barrier and to reach high rates in such cases, activating enzymes invest an extra input of energy that is not conserved and makes the activation highly irreversible. Oxygenases which involve a strong oxidant (O2/2 H2O, E0 = +0.818 V) and produce water besides the organic activation product are such energy-‘‘wasting’’ catalysts that achieve high rates. The reaction with methane is CH4 þ O2 þ NADH þ Hþ ! CH3 OH þ NADþ þ H2 O DG 0 ¼
344 kJ mol
1

ð11Þ

The sacrifice of energy to achieve activation via oxygenases is also reflected by the consumption of reducing equivalents detained from the energy-conserving respiratory chain: The oxygenase reaction consumes two reducing equivalents from the metabolism, and the insertion of the oxygen atom to yield the alcohol ‘‘cancels’’ two additional reducing equivalents; hence, four reducing equivalents are consumed. Dispite the significant amount of free energy dissipated and reducing power consumed by oxygenase reactions, this drain is not critical. The total free energy from the aerobic oxidation in this example is CH4 þ 2 O2 ! CO2 þ 2 H2 O DG  ¼
818 kJ ðmol CH4 Þ
1

ð12Þ

From the totally available 8 [H] per methane, 4 [H] are still available for the respiratory chain. With higher hydrocarbons, the drain of energy and reducing equivalents is even less relevant. An activation of hydrocarbons under anoxic conditions excludes oxygen11 and in the case of many catabolic net reactions with low energy gain strongly restricts the energy that can be dissipated to achieve activation. A reaction with particularly low net energy gain is the anaerobic oxidation of methane with sulfate (> Table 2). The activation reaction is most likely a reversal of the methyl-coenzyme M reductase (Mcr) reaction, the final step in methanogenesis which is exergonic under standard conditions (CoM
S
CH3 + HS
CoB ! CoM
S
S
CoB + CH4, DG =
30 [10] kJ mol
1; Shima and Thauer, 2005; Thauer and Shima, 2008). For methane activation, the standard free energy of the reverse Mcr (rMcr)

10

A prominent example is nitrogenase: Despite the long evolution of nitrogen fixation, an enzyme type has not evolved that catalyzes the thermodynamically feasible N2 reduction with H2 or energetically equivalent electron donors without an investment of energy. 11 The utilization of chlorate by facultatively anaerobic bacteria for hydrocarbon metabolism (Chackraborty and Coates, 2004; Tan et al., 2006) involves O2 that is generated from an intermediate (ClO2
! Cl
+ O2).

745

746

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

reaction in methane oxidizing archaea would thus be +30 [10] kJ mol
1. Methane activation with the disulfide CoM
S
S
CoB can therefore only take place if the products CoM
S
CH3 and HS
CoB are kept at very low concentration by effective scavenge in subsequent reactions. With respect to energy conservation in the total process, such a highly ‘‘concentration-controlled’’ reaction would be advantageous because it would be always very close to the equilibrium and not dissipate much energy. For the activation of the strong C
H-bond of methane (absolute value,
440 kJ mol
1; McMillen and Golden, 1982) by a thiyl radical or a NiIII center, which may be the most critical step, a decrease of the activation energy by a ‘‘dual stroke engine’’ mechanism was proposed (Thauer and Shima, 2008). rMcr has presumably two active sites, like Mcr. The release of the products from one site may transfer conformational energy to the other site where the substrates enter the reaction. However, this does not influence the equilibrium of the net activation reaction. Methane activation via a reversal of the Mcr reaction is not only of mechanistic but also of kinetic interest. The positive standard free energy change of the rMcr reaction sets severe limits to the rate of the formation of the initial intermediates. Using the Haldane equation,12 which connects the catalytic efficiencies of the forward and back reactions through an enzyme with the thermodynamic equilibrium constant of the reaction, the first step in AOM was estimated to be slower by a factor between 10
3 and 10
7 than the final step in methanogenesis (Shima and Thauer, 2005; Thauer and Shima, 2008). Also, the rate of the subsequent enzymatic step may be drastically limited by the low near-equilibrium concentrations of methylcoenzyme M and coenzyme B. The high content of the apparent rMcr in naturally enriched anaerobic methane oxidizers (Kru¨ger et al., 2003) may be a means to compensate for the slowness of the enzyme. The carbon–carbon addition of non-methane hydrocarbons at their methyl- or methylene group to fumarate is slightly exergonic (> Table 3) and to our present knowledge not associated with energy conservation. However, in view of the net energy gain with nonmethane hydrocarbons under anoxic conditions, such a loss is ‘‘affordable.’’ Only methane activation in an analogous way to yield methylsuccinate would be critical in an oxidation of methane with sulfate. The suggested mechanistic steps are an abstraction of a specific glycyl hydrogen in the polypeptide chain by a protein-activating enzyme (yielding
Gly▪
), subsequent hydrogen abstraction from a cysteyl group by the glycyl radical (yielding
CysS▪
), abstraction of a methyl hydrogen from toluene (yielding C6H5
▪CH2), addition of the benzyl radical to fumarate (yielding the benzylsuccinyl radical), and quenching of the radical to yield free benzylsuccinate and regenerate the cysteyl radical for the next catalytic round (chapter by Boll and Heider, this volume). Quantum chemical modeling of this reaction, for which a crystal structure of the enzyme was not available, supported the feasibility of the suggested steps (Himo, 2002, 2005). The rate-limiting step was calculated to be the addition of the benzyl radical to fumarate.

12

The Haldane equation describes the connection between the equilibrium concentrations of the reactants and products and their kinetic constants kcat and Km. The equilibrium constant is also thermodynamically given by the concentrations at DG = 0. In case of the reaction S ! P the connection is   ½P k S =KmS  ¼ cat ¼ e
DG =ðRT Þ : P =K P ½S eq kcat m

:

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

4

Quantitative Aspects of Cell Synthesis

4.1

ATP and Growth Yields

1

The more exergonic a catabolic reaction and the higher the efficiency of ATP synthesis (proportion of total free energy conserved in ATP), the more cell mass can be synthesized from a given substrate. The quantitative treatment of the efficiency of free energy conservation in the form of ATP and the amounts of cell mass formed with various substrates are subjects of an own area of research in microbiology. In this research, the measurable molar growth yield is of central interest, besides calculated free energy changes and ATP yields known from wellestablished pathways such as glycolysis. The molar growth yield, Y, is defined as the amount of cell dry mass, X (in g) per amount of totally consumed substrate, Stot (in mol). On the other hand, for indication of the energy gain from the catabolism, a growth yield with respect to the dissimiliated (viz. the energyyielding) proportion of the substrate, Sdiss, would be a more meaningful definition: Y ¼

X ðg mol
1 Þ Stot

Y diss ¼

X ðg mol
1 Þ Sdiss

ð13 a; bÞ

However, the latter definition and distinctive subscripts are not very common. Sdiss can be determined experimentally by quantifying the biomass, X, and the consumed electron acceptor (O2, NO3
, FeIII, or SO42
) or at least one of the products (CO2, N2, FeII/III, or H2S). The chemically formulated stoichiometric relationship between substrate and product (> Table 1) then reveals Sdiss, which leads to Ydiss (13b). The fraction of the dissimilated substrate as part of the totally consumed substrate in anaerobic bacteria is usually much higher than in aerobic bacteria:     Sdiss Sdiss > ð14Þ Stot anaerobic Stot aerobic Some measured growth yields of aerobic and anaerobic hydrocarbon utilizing microorganisms are listed in > Table 4. If consumption of the electron acceptor or formation of the catabolic product has not been quantified, or if only a Y value (13a) has been reported, Ydiss can be calculated. With Sass for the assimilated amount of substrate, the totally consumed substrate is Stot ¼ Sdiss þ Sass

ð15Þ

Division by the obtained cell mass yields Stot Sdiss Sass ¼ þ X X X

ð16Þ

1 1 Sass þ ¼ X Y Ydiss

ð17Þ

and with definitions (13a, b)

The expression Sass/X may be termed the assimilatory substrate demand, dass (in mol g
1). The reciprocal term X/Sass can be defined as another type of yield, the amount of cell mass (in g) obtained per assimilated amount of substrate (in mol), and designated Yass. The connection is thus dass = 1/Yass. This leads to

747

Toluene, O2

Naphthalene, O2

Pseudomonas putida

Pseudomonas sp.

Toluene, SO42


Desulfobacula toluolica 0.31

0.059

0.037

0.83

0.50*

0.70*

1.0*

1.28*

1.20*

0.50

0.9*

29*

13.5*g

0.59

88*

64.1

64.5

92.1

100

94.7

120*g

401

226

204

127f to 280f

125

molar, Y (g mol
1)

33*

13.8

0.6*

129*

82

91.6

159

185

197

148

1240

366

310

164f to 563f

214

molarc, Ydiss (g mol
1)

0.88 (88%)

0.97 (97%)

0.99 (99%)

0.68 (68%)

0.78 (78%)

0.70 (70%)

0.58 (58%)

0.54 (54%)

0.48 (48%)

0.81 (81%)

0.32 (32%)

0.62 (62%)

0.66 (66%)

0.77 (77%) to 0.50 (50%)

0.58 (58%)

Rabus et al. (1993)

So and Young (1999)

Nauhaus et al. (2007)

Rabus and Widdel (1995)

Wodzinski and Johnson (1968)

Dinkla et al. (2001)

Bordel et al. (2007)

Reardon et al. (2000)

Reardon et al. (2000)

Bonin et al. (1992)

This article (see text)

Einsele (1983)

Einsele (1983)

Wagner et al. (1969)

Wodzinski and Johnson (1968)

Reference

b

a

Only directly measured (‘‘real’’) growth yields are listed and not Ymax values obtained from extrapolation Original value from the reference is indicated by asterisk; other values were calculated for this chapter (see text). Ydiss wass calculated via (19). The needed Yass was calculated according to > Table 3, assuming the biomass bulk formula C4H7O3; the Yass values (g mol
1) are as follows: methane, 48.5; n-octane, 303; n-pentadecane, 558; n-hexadecane, 594; n-heptadecane, 630; benzene, 182; toluene, 218; ethylbenzene, 255 c Relative to dissimilated substrate d Calculated according to Sdiss/Stot = Y/Ydiss e Additional assimilation of added yeast extract is likely f For convenience calculated with pentadecane (which was part of the mixture) g With 2% O2 in gas phase; with more O2 the yield decreased h Estimated by assuming that 55% of cell dry mass is protein

n-Hexadecane, SO42


Deltaproteobacterium, strain AK-01

Archaea (ANME-2) + Deltaproteobacteria Methane, SO42


Betaproteobacterium, strain EbN1

Ethylbenzene, NO3


Toluene, O2

Pseudomonas putida

Anaerobic

Benzene, O2

n-Heptadecane, O2

Pseudomonas nautica

Toluene, O2

n-Hexadecane, O2

Ideal aerobe

Pseudomonas putida

1.77

n-Hexadecane, O2

Candida tropicalis

Pseudomonas putida

1.0*

n-Hexadecane, O2

Micrococcus cereficans

0.60*e to 1.32*

n-Octane, O2

Long-chain n-alkane mixture, O2

Pseudomonas sp. 1.1*

by mass (g g
1)

Nocardia sp.

Aerobic

Hydrocarbon and electron acceptor

Fraction of substrate catabolizedd, Sdiss/Stot

1

Microorganism

Growth yieldb (Cell dry mass per amount of hydrocarbon)

. Table 4 A selection of growth yieldsa of aerobic and anaerobic microorganisms on hydrocarbons, and calculated fraction of the dissimilated substrate

748 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1

1 1 1 þ ¼ Y Ydiss Yass

ð18Þ

Yass  Y Yass
Y

ð19Þ

Rearrangement leads to Ydiss ¼

The values for dass or Yass are calculated from chemically formulated stoichiometries. This requires the assumption of bulk formulas for cell dry mass. The simplest bulk formula is that of carbohydrates, hCH2Oi. For aerobic methanotrophs, the formula hC4H8O2Ni was used (van Dijken and Harder, 1975). A simpler N-free variant with the same bulk oxidation state of carbon is hC4H7O3i (Pfennig and Biebl, 1976). A precise yet more complicated formula, hC4.36H8.24O1.87Ni, was determined for an aerobic bacterium grown with heptadecane (Bonin et al., 1992). Considering the oxidation state of carbon is more important than including nitrogen. Because in the case of hydrocarbons the substrate carbon is more reduced than cell mass carbon, CO2 is included in the assimilation equations (> Table 1). Now, also the fraction of the dissimilated substrate as part of the totally consumed substrate can be calculated even if only a Y value is available from the literature: Sdiss Yass
Y ¼ Stot Yass

ð20Þ

For instance, for aerobic growth with hexadecane (M = 226.45) a growth yield by mass of 1 g (g C16H34)
1 has been reported, which equals a molar growth yield of Y = 226 g (mol C16H34)
1. The assimilation equation is 17 C16 H34 þ 120 CO2 þ 54 H2 O ! 98 C4 H7 O3

ð21Þ

dass ¼ 1:68  10
3 mol g
1 ; Yass ¼ 594 g mol
1 Equation (19) yields Ydiss = 366 g mol
1. The fraction of the dissimilated substrate is Sdiss ¼ 0:62 ðor 62%Þ Stot

ð22Þ

Above all, Y values are expected to provide information about the ATP yield as a parameter of high relevance to understand the efficiency of or losses in the energy flow Free energy of catabolic reaction # Free energy in formed ATP # Free energy ðor ATPÞ consumed for cell synthesis The ATP yield or qATP, is the amount of ATP (in mol) formed per amount of dissimilated substrate (in mol). At first glance, the concept appears straight-forward. From anaerobic pathways with biochemically known qATP , as for instance the homolactic fermentation of glucose (qATP = 2), the amount of cell mass obtained per mol ATP, the so-called YATP , can be calculated from the determined growth yield via YATP = Ydiss /qATP . If for another bacterium of interest the qATP is unknown but Ydiss has been determined, this should in principle allow to

749

750

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

calculate the desired qATP parameter via qATP = Ydiss /YATP.13 However, there is a serious drawback in that determined YATP values, viz. the energy expenses for biomass synthesis, vary enormously for different growth substrates and among various bacteria. This is not surprising because synthesis of an amount of biomass for instance from free acetate as the growth substrate needs more ATP than synthesis from carbohydrates and amino acids added to the medium. But even with the same substrate for biosynthesis, determined YATP values among bacteria vary significantly. These problems are treated by the calculation of theoretical ATP demands for the synthesis of biomass with its diverse fractions (polysaccharides, protein, etc.) from starting substrates, and by consideration of the fractions of energy or ATP that do not lead to productive growth. This non-productive consumption of energy or ATP is interpreted as maintenance energy (Pirt, 1965; Tempest and Neijsesel, 1984), an uncoupling of the anabolism from the catabolism at varying extent, or an extra ‘‘spill’’ of energy (Russell, 2007) in addition to the ‘‘regular’’ dissipation. In the concept of Pirt (1965), the proportion of the substrate consumed per time for maintenance rather than for productive growth is regarded as a constant that is independent of the growth rate, m. Hence, the slower the growth of a bacterium and the lower the biomass production per time, the higher the proportion of the substrate consumed for maintenance. If therefore growth yields of an organism at different growth rates are extrapolated to a theoretical infinitely high growth rate (no time required for growth) in a plot of 1/Ydiss versus 1/m, the proportion of the substrate consumed for maintenance should become zero. At 1/m = 0 (m = 1) the theoretically highest growth yield, Y max (more precisely Ydissmax) is obtained that is used to gain information about qATP and YATP. Such concepts have been applied to vast series of non-hydrocarbon substrates (Heijnen and van Dijken, 1992; Stouthamer 1988). In the case of hydrocarbons, aerobic methanotrophs (Leak and Dalton, 1985; van Dijken and Harder, 1975) and degraders of long-chain alkanes (Erickson, 1981; Ferrer and Erickson, 1979) have been of interest for such mainly theoretical studies. If the catabolism of a substrate is likely to involve conventional biochemical reactions (-oxidation, citric acid cycle, dehydrogenations with NAD+ and flavoenzymes, etc.) and an aerobic respiratory chain, a qATP value can be also predicted from the ATP-yielding reactions. Via YATP values determined in other studies, a Y max can be subsequently predicted and compared to an experimentally determined one. As an example, > Fig. 5 presents the catabolic scheme for aerobic degradation of hexadecane with qATP = 124 (mol/mol). According to the free energy change of the reaction (
10 392 kJ mol
1; > Fig. 5), the average energy need for ATP synthesis would be 100 kJ (mol ATP)
1. If a YATP of 10 g cell dry mass (mol ATP)
1 is assumed that is likely for cell synthesis from the hexadecane-derived acetate units (Erickson, 1981; Stouthamer, 1988), this would lead to Ydiss = 1240 g cell mass (mol C16H34)
1. The Y value is obtained via a transformation of (19): Y ¼

Yass  Ydiss Yass þ Ydiss

ð23Þ

This yields (with the above Yass = 594 g mol
1) a value of Y = 401 g mol
1, which would be a yield by mass of 1.77 g cell mass (g C16H34)
1. This may be regarded as an ‘‘ideal’’ yield with hexadecane. The fraction of dissimilated hexadecane would be only The qATP is conceptually related to the P/2e
ratio in aerobic and anaerobic respiration which indicates the number of ATP molecules formed per electron pair transported in the respiratory chain (in aerobes also P/O ratio). However, the qATP also includes ATP from substrate level phosphorylation.

13

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1

. Figure 5 Reducing equivalents and ATP synthesis in the aerobic catabolism of hexadecane (C16H32, M = 226.45). Reducing equivalents from enzyme-bound FADH2 enter the respiratory chain at the quinone (Q) level. The assumed proton translocation in the respiratory chain underlying this scheme is 10 H+/NADH and 6 H+/QH2. A phosphorylation yield of 1 ATP per 3.5 H+ was arbitrarily assumed here (based on the commonly assumed range of 1 ATP per 3 to 4 H+). The resulting net ATP yield is thus 124 mol ATP per mol C16H34, 0.55 mol ATP per g C16H34, or 5.3 mol ATP per mol O2. For comparison: Glucose (C6H12O6, M = 180.16) would yield 10 NADH and 2 QH2 allowing formation of 32 ATP via respiration; with 4 ATP from glycolysis and the tricarboxylic acid cycle the net yield is 36 mol ATP per mol C6H12O6, 0.20 mol ATP per g C6H12O6, or 6.0 mol ATP per mol O2.

Sdiss ¼ 0:32 ðor 32%Þ Stot

ð24Þ

most of the substrate is therefore assimilated. The lower yields from experiments (> Table 4) indicate significant energy consumption for maintenance or by uncoupling.

4.2

Requirement for Minerals (N, P, Fe)

Growth yields are not only of basic but also of practical interest because they can be used to estimate the amount of essential minerals required for oil-degrading bacteria. Since crude oil has an extremely low content of nitrogen, phosphorous and iron, these important elements are often the limiting ones in oil biodegradation. Availability of sulfur is usually not a problem, because oil contains organic sulfur and many natural waters are rich in sulfate (seawater: 28 mM).

751

752

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

In the environment and in cultures, microorganisms often obtain the limiting elements as inorganic species (NH4+, NO3
, H2PO4
/HPO42
, Fe2+, FeIII minerals, etc.). The above bulk formula for cell mass which considers nitrogen, hC4H8O2Ni, suggests a content of 14% N by mass; it does not consider phosphorus and iron. The extended Redfield ratio, (CH2O)106(NH3)16(H3PO4) = hC106H263O110N16Pi, which was derived from the originally determined molar C:N:P ratio of 106:16:1 of marine phytoplankton (Brewer et al., 1997), considers in addition phosphorus. Carbon in this formula has the bulk oxidation state as in carbohydrates, which may not very precisely reflect bacterial cell mass. ‘‘Redfield biomass’’ contains 6.3% N and 0.9% P by mass. With these ratios, 1 g biomass produced aerobically during complete consumption of 1 g (1.3 ml) hexadecane would need 0.24 g (4.5  10
3 mol) NH4Cl and 0.04 g (0.3  10
3 mol) KH2PO4. In a marine environment with for instance 1 mM combined nitrogen and 0.06 mM phosphate, the microbial cell mass produced with 1 g hexadecane would consume the nitrogen and phosphorous from roughly 5 m3 water. However, such calculations should be applied reservedly in the study of natural hydrocarbon bioremediation. A lower in situ growth yield and N and P release from lysed cells may result in a lower than the calculated need for N and P. On the other hand, oil as a hydrophobic substrate is not distributed like soluble organic carbon in the water body but forms buoyant layers. Cells of hydrocarbon-degrading bacteria largely depend on physical contact with the oil, so that supply of biominerals by advective transport is a severely limiting factor (chapter by Harms et al. in this volume). The controlled use of environmentally friendly immobilized N and P sources (as well as of iron sources that have not been considered here) that tend to stay in contact with oil may therefore be a justified method to stimulate oil degradation in eutrophic waters (chapter by Ron and Rosenberg in this volume).

5

Research Needs

The application of thermodynamic data to microbial systems as a whole is a theoretical approach that is basic for the understanding of the overall catabolism of chemotrophic microorganisms (Thauer et al., 1977). Even though it is not regarded as an own field of microbiological research, the underlying formalism accompanies the study of numerous metabolic types of bacteria and may lead to the recognition of scientifically challenging questions that have not been encountered before. One of these is clearly the appropriate understanding of how microorganisms conserve energy at low chemical potential, viz. with low-energy substrates and combinations of electron donors and electron acceptors with marginal differences in their redox potential. Prominent processes of such type are anaerobic reactions involving hydrocarbons, such as the anaerobic oxidation of methane or conversion of non-methane hydrocarbons to methane by microbial consortia which even have to share the low net energy gain. Also individual enzymatic reactions in the anaerobic degradation of hydrocarbons, in particular the activating steps and intermediate energetic states (energy-rich transition states) need a deeper understanding from an energetic and kinetic point of view. There may be even open questions concerning growth yields and the efficiency of energy conservation during growth with hydrocarbons under various environmental conditions. Their examination could be relevant for the study of hydrocarbon bioremediation in oligotrophic aquatic environments.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1

Appendix . Table 5 Hydrocarbons (methane, propane, n-hexane and benzene as examples) and other substances as ‘‘energy carriers.’’ In a reaction with oxygen, liquid hydrocarbons reveal a high gravimetric energy density in comparison to many other compounds and elements (calculated for the highest oxides in their standard state). Gaseous hydrocarbons reveal a high volumetric energy density Substance

DG of oxidiation with O2

DH of oxidiation with O2

Per mass of substance (kJ kg
1)

Per volume of substance (kJ m
3)

Per mass of substance (kJ kg
1)

Per volume of substance (kJ m
3)

H2


117.6  103


9.7  103


141.8  103


11.7  103

CH4


51.0  10


55.5  10


36.4  103

Gases (101 kPa)

C3H8 NH3

b

3


33.5  10

3

3


47.8  103


86.2  103 a


50.3  103


90.8  103 a


19.9  10


13.8  10

3


22.5  10


15.6  103 a

3

3 a


19.3  103


26.8  103 a


23.3  103


32.4  103 a

Li


40.4  103


21.6  106


43.0  103


23.0  106

B


55.2  103


135.7  106


58.8  103


144.7  106

CGraphite


32.8  10

3


74.4  10

6


32.8  10


74.4  106

C6H14


46.7  10

3


30.8  10

6

3


48.3  10


31.9  106

C6H6


41.0  103


36.0  106


41.8  103


37.5  106

CH3OH


21.9  10

3


17.5  10

6


22.7  10


18.1  106

CH3CH2OH


28.8  10

3


22.8  10

6

3


29.7  10


23.5  106

C6H12O6 (a-D-Glucose)


16.0  103


25.0  106


15.6  103


24.3  106

Mg


23.4  103


40.8  106


24.8  103


43.1  106

Al


29.3  10


79.2  10

3


31.1  10


84.0  106

Si


30.5  103


71.1  106


32.4  103


75.5  106

Pwhite


21.8  10

3


39.7  10

6


24.1  10


43.9  106

S


11.6  10

3


22.7  10

6

3


12.3  10


24.1  106


7.4  103


58.3  106

H2S c Solids or liquids

Fe a

3


6.6  103

6


52.3  106

3

3

3

For convenience, ideal behavior assumed. In reality, the volumetric energy density will be somewhat higher If N2 (g) is produced c If H2SO4 (l) is produced b

753

754

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

. Table 6 Thermodynamic properties of hydrocarbons and other compounds. Data are from other compilationsa Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Formula mass (g mol
1)

Free energy of Enthalpy of formation from the formation from the elements, DGf elements, DHf (kJ mol
1) (kJ mol
1)

Entropyb, S˚ (J K
1 mol
1)

Alkanes
74.81c
74.9d

186.26c 186.2e


32.82c
32.6e
32.89f


84.68c
83.8d

229.60c 229.1d

44.096


23.49c
23.6d
23.4e


103.85c
104.7d

269.91c 270.2d

n-Butane (g)

58.123


17.03c
17.2d
15.7e


126.15c
125.6d
125e

310.23c 310.1d

2-Methylpropane (g)

58.123


20.9 d
18.0e


134.2 d
132e

294.6d 295e

n-Pentane (l)

72.150


9.3d
9.21e


173.1c
173.5d

262.7d

2-Methylbutane (l)

72.150


179e

260e

Methane (g)

16.043


50.72c
50.8d
50.75f

Methane (aq)

16.043


34.4g

Ethane (g)

30.069

Propane (g)


14.6e

n-Hexane (l)

86.177


3.8
4.28e


198.7
198.8d
199e

296.1d 296e

2-Methylpentane (l)

86.177


8.11e


204e

291e

d

c

n-Heptane

100.203

1.0 1.28e


224.4

328.6c 328e

n-Octane (l)

114.23

6.41e


249.9c

361.1c

114.23

3.85

e


255.1

352e

e


252

c

358e


252c

350e

2-Methylheptane (l)

c

c

c

3-Methylheptane (l)

114.23

4.68

4-Methylheptane (l)

114.23

7.8e

n-Decane (l)

142.28

17.5d 17.4e


300.9d

425.5d 426e

n-Dodecane (l)

170.34

28.1d 28.4e


350.9d
352e

490.6d

n-Tridecane (l)

184.36

33.8e


378e

523e

n-Tetradecane (l)

198.39

38.8e


403e

555e

n-Hexadecane (l)

226.44

49.8h 52.2i


454.4h

n-Heptadecane (l)

240.47

n-Octadecane (c)

254.50


480e 53.9e


569e

497e

Cyclopentane (l)

70.134

36.4 36.5e


105.1
106e

204.3d

Cyclohexane (l)

84.161

26.8e 26.7d


156c
156.4d

204.4d

d

d

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization . Table 6 (Continued) Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Formula mass (g mol
1)

Free energy of Enthalpy of formation from the formation from the elements, DGf elements, DHf (kJ mol
1) (kJ mol
1)

Entropyb, S˚ (J K
1 mol
1)

Unsaturated hydrocarbons, non-aromatic Ethene (g)

28.054

68.15c 68.4d

52.26c 52.5d

219.56c 219.3d

Propene (g)

42.080

62.78c 62.8d 74.8e

20.42c 20.0d 20.4e

267.05c 266.6d 227e

1-Butene (g)

56.107

71.39c 71.3d 72.0e


0.13c 0.1d 1.17e

305.71c 305.6d 307e

cis-2-Butene (g)

56.107

65.95c 65.9d 67.3e


6.99c
7.1d
5.70e

300.94c 300.8d

trans-2-Butene (g)

56.107

63.06c 63d 64.3e


11.17c
11.4d
10.1e

296.59c 296.5d 296e

Ethyne (g)

26.038

209.2c

226.73c

200.94c

Benzene (l)

78.113

124.3c 124.4d 124.5f

49.0c

173.3c 173.4d

Toluene (l)

92.140

113.8d 110e 114.22f

12.4d 8.08e

221d 219e

Aromatic hydrocarbons

Ethylbenzene (l)

106.17

120e


12.5e

255e

e

246.5d 246e

o-Xylene (l)

106.17

110.3 111e


24.4

m-Xylene (l)

106.17

107.7d


25.4e

252.2d

p-Xylene (l)

106.17

d

110.1

d


24.4
24.3e

247.4d

1,3,5-Trimethylbenzene (l)

120.19

103.9d


63.4d
63.5e

273.6d 273e

Naphthalene (c)

128.17

201d

78.53c 77.9d

166.9d

1-Methylnaphthalene (l)

142.20

189.4d

56.3d

254.8d

2-Methylnaphthalene (c)

142.20

192.6d

44.9d

220d

154.21

d

d

205.9d

d

207.6d 207e

Biphenyl (c)

d

254.2

d

99.4

Anthracene (c)

178.23

286.0

129.2 128e

Phenanthrene (c)

178.23

268.3d

116.2d 113e

211.7d 212e


238.66c
239.1d

126.8c 127.2d


277.69c
277.0d

160.7c 161.0d

Alcohols, phenolic compounds Methanol (l)

32.042


166.27c
166.8d

Methanol (aq)

32.042


175.39f

Ethanol (l)

46.069


174.78c
174.2d

755

756

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

. Table 6 (Continued) Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Formula mass (g mol
1)

Free energy of Enthalpy of formation from the formation from the elements, DGf elements, DHf (kJ mol
1) (kJ mol
1)

Ethanol (aq)

46.069


181.75f

1-Propanol (l)

60.096


170.6d

1-Propanol (aq)

60.096


175.81

2-Propanol (l)

60.096


180.3d
182e

2-Propanol (aq)

60.096


185.94f

1-Butanol (l)

74.122


162.5d

1-Butanol (aq)

74.122


171.84f

2-Butanol (l)

74.122

Entropyb, S˚ (J K
1 mol
1)


302.6d

194.6d


318.1d
319e

180.6d 180e


327.3d

226.4d 252e


177.0d


342.6d

225.1d

d

f

1-Hexadecanol (c)

242.44


98.7
98.8e


686.7
684e

451.9d 452e

Benzyl alcohol (l)

108.14


27.5d
27.3e


160.7d

216.7d


50.9c
50.4d
47.5e
47.6f


165c
165.1d
163e

146c 144d 142e


210.0d


361.1d

150.2d


108.57c
116e

218.77c


192.30c
191.8d

160.2c 160.4d


247e

247e

Phenol (s)

1,2-Dihydroxybenzene (s)

94.113

124.14

d

Aldehydes, ketones Formaldehyde (g)

30.026


102.53c
109.9d
111e
112.97f

Formaldehyde (aq)

30.026


130.54f

Acetaldehyde (l)

44.053


128.12c
128.3d

Acetaldehyde (aq)

44.053


139.9f

Butyraldehyde (l)


127e

Acetone (l)

58.080


155.4
155.8d

Acetone (aq)

58.080


161.17f

2-Butanone (l)

72.107

Benzaldehyde (l)

106.12

c


248.1
242.1d

200.4c 200.6d


151.4d
156e


273.3d
279e

238.8d 241e

9.4d


87.0d

c

Carboxylic acids, carboxylates Formic acid (l)

46.026


361.35c
360e


424.72c
425.1d
423e

128.95c

Formic acid (aq)

46.026


356.3j


410.3j

163.7j

Formate (aq)

45.018


351.04
334.9j


410.3

91.7 j

Acetic acid (l)

60.052


389.9c
390.2d
392e


484.5c
484.4d

159.8c 159.9d

Acetic acid (aq)

60.052


396.46c


485.76c

178.7c

59.045


369.31
369.41f


486.01

86.6c







Acetate (aq)

f

c

j

c

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization . Table 6 (Continued) Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous)

Formula mass (g mol
1)

Free energy of Enthalpy of formation from the formation from the elements, DGf elements, DHf (kJ mol
1) (kJ mol
1)

Propionate
(aq)

73.071


361.08f

Butyrate
(aq)

87.098


352.63f




Hexanoate (aq)

115.15


335.96f

Palmitic acid (c)

256.43


305.0f


882e

Benzoic acid (c)

122.12


245.3
245.6f

Benzoate
(aq)

121.12


229.3k

Fumarate

c

2


(aq)

114.06


604.21f

2


(aq)

116.07


690.23f

Succinate

130.10


681.6 to
685.5l

(1-Methylpentyl)succinate2
(aq)

200.23


644.0 to
647.3l

Benzylsuccinate2
(aq)

206.20


521.1 to
525.4l

Methylsuccinate

2


(aq)

Entropyb, S˚ (J K
1 mol
1)

452e c


385.1
385.2d

167.6c


23.0d
28e

242.6d 242e

Hydrocarbon-derived nitrogen-, oxygen-, sulfur-, and halo-compounds 31.057

32.3d 27.5e

Methylammonium+ (aq)

32.065


40.0f

Ethylamine (g)

45.084

37.3d


47.4d
48.5e

289.9d

Pyridine (l)

79.101

181.3d

100.2d

177.9d

Pyridine (aq)

79.101

177.1f

Aniline (l)

93.128

149.2d 148e

31.3d 29.7e

191.4d 192e

Dimethylether (g)

46.069


112.9d
114e


184.1d
185e

267.1d

Diethylether (l)

74.122


116.7d


279.3d

253.1d

Methylamine (g)

Methanethiol (g)

48.10


9.9 0.754e


22.9
12.4e

255.1d

Dimethylsulfide (l)

62.13

5.72e


65.4e

196e


73.7

e

207e

d

Ethanethiol (l)

62.13

d


5.7

e d

d

Thiophene (l)

84.14

121.2 122e

80.6 81.7e

181.2d

Thiophenol (l)

110.17

134d

64.1d 62.8e

222.8d

Fluoromethane (g)

34.033


213.8d
222e


237.8d
247e

222.8d

Tetrafluoromethane (g)

88.005


888.3d
862e


933.6d
908e

261.3d 262e

Chloromethane (g)

50.488


58.5d
58.1e


81.9d
82.0e

234.2d 233e

96.944


63.3e


117e

179e

Trichloromethane (l)

119.38


71.2e


132e

203e

Tetrachloromethane (l)

153.82


62.6d
68.4e


132.8d
139e

216.2d 214e

Dichloromethane (l)

Chloroethane (g)

64.515


60.5d
53.0e


112.1d
105e

275.8d 275e

Fluorobenzene (l)

96.104


69.0e


145e

206e

757

758

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

. Table 6 (Continued) Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous) Chlorobenzene(l)

Formula mass (g mol
1) 112.56

Free energy of Enthalpy of formation from the formation from the elements, DGf elements, DHf (kJ mol
1) (kJ mol
1) 89.2d 93.7e

11.0d 10.6e

Entropyb, S˚ (J K
1 mol
1) 209.2d 194e

Inorganic compounds H2 (g)

2.0158

0

0

H+ (aq), pH = 0

1.0074

0

0

H+ (aq), pH = 7

1.0074

C, graphite (c)

12.011

(DGf0 )
39.97l 0

130.684c 0 (DS0 ) 134.06

0

5.740c

0 c

c

12.011

2.900

CO (g)

28.011


137.17c
137.15f


110.53c

197.67c

CO2 (g)

44.010


394.36c
394.39d


393.51c
393.52d

213.74c 213.80d

CO2 (aq)

44.010


385.98c
386.02f


413.80c

117.6c

HCO3
(aq)

61.017


586.77c
586.85f


691.99c

91.2c

CO32
(aq)

60.009


527.81c
527.90f


677.14c


56.9c

N2 (g)

28.0134

0

0

191.61c

NH4+ (aq)

18.038


79.31c
79.37e


132.51c

113.4c

N2O (g)

44.013

104.20c 104.18f

82.05c

NO2
(aq)

46.006


37.2f,m
34.54j


106.3j
104.6m

125.2j 140m

NO3
(aq)

62.005


108.74c
111.34f
110.7j


205.0c
206.7j
207.3m

146.4c 146.5j

O2 (g)

31.999

0

0

205.138c 205.147d

H2O (l)

18.015


237.13c
237.14d
237.178f
237.18m


285.83c

69.91c 69.95d

S, (a, rhombic; c)

32.06

0

0

31.80c 32.056d

H2S (g)

34.08


33.56c
33.3d


20.63c
20.5d

205.79c 205.7d

H2S (aq)

34.08


27.83c
27.87d


39.7c
39.8d

121c

HS
(aq)

33.072

12.08c 12.05m


17.6c

62.08c

SO42
(aq)

96.06


744.53c
744.63f


909.27c

20.1c

F
(aq)

18.999


278.79c


332.63c


13.8c

35.453


131.23
131.3m


167.16




Cl (aq)

c

1.895 1.897d

2.377c

C, diamond (c)

c

219.85c

56.5c

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization . Table 6 (Continued) Compound (Standard states: g, gaseous; l, liquid; c, crystalline; aq, aqueous) Mn2+ (aq) MnCO3

Formula mass (g mol
1) 54.937 114.95

Free energy of Enthalpy of formation from the formation from the elements, DGf elements, DHf (kJ mol
1) (kJ mol
1)

Entropyb, S˚ (J K
1 mol
1)


227.8j
228.0m


223.3j
220.7m


84j
73.6m


816.0m


889.3m

100m


465.1m


520.0m

MnO2

86.937

Fe2+ (aq)

55.846


78.90c
78.87m

Fe3+ (aq)

55.845


4.7c
4.60m

53m


89.10m


137.7c


48.5c


315.9c

FeCO3 (siderite; c)

115.86


674.3j
666.7m


748.2j
737.0m

92.9j 105m

Fe(OH)3 (amorphous)

106.87


695j
699m


824.8j

96j

Fe2O3 (a, hematite; c)

159.69


742.2c
742.7m


824.2c
824.6m

87.40c

Fe3O4 (magnetite; c)

231.54


1015.4c
1012.6m


1118.4c
1115.7m

146.4c

a Original sources cited in the used compilations were not consulted. If a precise and rounded value is given in the compilations, the precise value is indicated here b The absolute entropy values may be used to calculate entropy changes of reactions as well as entropies of formation, DSf ; the latter can be used to prove consistency of literature data via DGf = DHf
298.15 DSf . Example of n-hexane (C6H14): DSf = 291.6
(6  5.74)
(7  130.68) =
657.6 J K
1 mol
1. DGf =
198.8
(298.15 
0.6576) =
2.7 kJ mol
1. The result is close to the value given in the literature source (
3.8 kJ mol
1) c Atkins and de Paula (2006) d Dean (2004) e D’Ans and Lax (1983) f Thauer et al. (1977) g Calculated via solubility of 1.39 mol l
1 atm
1 at 25  C (from Wilhelm et al., 1977) h Via extrapolation or interpolation of the listed data i Zengler et al. (1999) j Garrels and Christ (1965) (data transformed by using 1 cal = 4.1868 J) k Widdel et al. (2007) l Free energy associated with dilution of 1 mol H+ from a = 1 to a = 10
7, which is R T ln 10
7 m Stumm and Morgan (1981)

759

1

. Figure 6 The two standard states (framed) of methane. Aqueous methane, CH4 (aq), in its standard state which corresponds to a very high partial pressure has a higher energy content than gaseous methane, CH4 (g), in its standard state. Hence, indication of the DG oder DG0 of a formulated reaction involving methane must indicate the applied standard state. Calculation of the free energy change of a reaction for real (measured) pressures or concentrations (according to (4)) must yield the same result with each standard state. Application of the gaseous standard state for calculation is also justified if there is no gas phase. Most natural conditions will be closest to the gaseous standard state. The DGf of CH4 (aq) was calculated via the solubility of 0.00139 mol l
1 atm
1 (Wilhelm et al., 1977), assuming that this concentration is numerically equivalent with the activity of CH4 (aq) that is in equilibrium with CH4 (g) of standard pressure. In seawater, the dissolved methane concentration in equilibrium with gaseous methane of standard pressure is lower (Yamamoto et al., 1976), even though this has the same activity as methane in pure water.

760 Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

. Figure 7 The three standard states (framed) of inorganic carbon (CO32
not included), the product of hydrocarbon oxidation. Most natural conditions will be closest to the gaseous standard state. See also remarks in legend of Appendix > Fig. 6. Reactions are indicated for pH = 7.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

1 761

762

1

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization

References Anderson RT, Lovley DR (2000) Hexadecane decay by methanogenesis. Nature 404: 722–723. Atkins PW, de Paula J (2006) Physical chemistry, 8th edn. Oxford: Oxford University Press. Bonin P, Gilewicz M, Bertrand JC (1992) Efects of oxygen on Pseudomonas nautica grown on n-alkane with or without nitrate. Arch Microbiol 157: 538–545. Bordel S, Mun˜oz R, Dı´az L, Villaverde S (2007) New insights on toluene biodegradation by Pseudomonas putida F1: influence of pollutant concentration and excreted metabolites. Appl Microbiol Biotechnol 74: 857–866. Brewer PG, Goyet C, Friedrich G (1997) Direct observation of the oceanic CO2 increase revisited. Proc Nat Acad Sci USA 94: 8308–8313. Chakraborty R, Coates JD (2004) Anaerobic degradation of monoaromatic hydrocarbons. Appl Microbiol Biotechnol 64: 437–446. D’Ans J, Lax E (1983) Taschenbuch fu¨r Chemiker und Physiker, Bd 2, 2. Berlin: Aufl. Springer. Dean JA (2004) Lange’s handbook of chemistry, 16th edn. New York: McGraw-Hill. Dinkla IJT, Gabor E, Janssen DB (2001) Effects of iron limitation on the degradation of toluene by Pseudomonas strains carrying the TOL (pWWO) plasmid. Appl Environ Microbiol 67: 3406–3412. Dolfing J, Larter SR, Head IM (2008) Thermodynamic constraints on methanogenic crude oil biodegradation. ISME J 2: 442–452. Einsele A (1983) Biomass from higher n-alkanes. In H-J Rehm, G Reed (eds.) Biotechnology, vol 3. Weinheim: Verlag Chemie, pp. 43–81. Erickson LE (1981) Energetic yields associated with hydrocarbon fermentations. Biotechnol Bioeng 23: 793–803. Ettwig KF, Shima S, van de Pas-Schoonen KT, Kahnt J, Medema MH, Op den Camp HJ, Jetten MS, Strous M. (2008) Denitrifying bacteria anaerobically oxidize methane in the absence of Archaea. Environ Microbiol 10: 3164–3173. Ferrer A, Erickson LE (1979) Evaluation of data consistency and estimation of yield parameters in hydrocarbon fermentations. Biotechnol Bioeng 21: 2203–2233. Garrels RM, Christ CL (1965) Solutions, minerals and equilibria. New York: Harper & Row. Heijnen JJ, Van Dijken JP (1992) In search of a thermodynamic description of biomass yields for the chemotrophic growth of microorganisms. Biotechnol Bioeng 39: 833–858. Himo F (2002) Catalytic mechanism of benzylsuccinate synthase, a theoretical study. J Phys Chem B 106: 7688–7692.

Himo F (2005) C
C bond formation and cleavage in radical enzymes, a theoretical perspective. Biochim Biophys Acta 1707:24–33. Jackson BE, McInerney MJ (2002) Anaerobic microbial metabolism can proceed close to thermodynamic limits. Nature 415: 454–456. Jones DM, Head IM, Gray ND, Adams JJ, Rowan AK, Aitken CM, Bennett B, Huang H, Brown A, Bowler BF, Oldenburg T, Erdmann M, Larter SR (2008) Crude-oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 451: 176–180. Kru¨ger M, Meyerdierks A, Glo¨ckner FO, Amann R, Widdel F, Kube M, Reinhardt R, Kahnt J, Thauer RK, Shima S (2003) A conspicuous nickel protein in microbial mats that oxidise methane anaerobically. Nature 426: 878–881. Leak DJ, Dalton H (1985) Growth yields of methanotrophs. Appl Microbiol Biotechnol 23: 477–481. McMillen DF, Golden DM (1982) Hydrocarbon bond dissociation energies. Ann Rev Phys Chem 33: 493–532. Nauhaus K, Boetius A, Kru¨ger M, Widdel F (2002) In vitro demonstration of anaerobic oxidation of methane coupled to sulphate reduction in sediment from a marine gashydrate area. Environ Microbiol 4: 296–305. Nauhaus K, Albrecht M, Elvert M, Boetius A, Widdel F (2007) In vitro cell growth of marine archaealbacterial consortia during anaerobic oxidation of methane with sulphate. Einviron Microbiol 9: 187–196. Pirt SJ (1965) The maintenance energy of bacteria in growing cultures. Proc Royal Soc London 163B: 224–231. Pfennig N, Biebl H (1976) Desulfuromonas acetoxidans gen. nov. and sp. nov., an new anaerobic, sulfurreducing, acetate-oxidizing bacterium. Arch Microbiol 110: 3–12. Rabus R, Widdel F (1995) Anaerobic degradation of ethylbenzene and other aromatic hydrocarbons by new denitrifying bacteria. Arch Microbiol 163: 96–103. Rabus R, Nordhaus R, Ludwig W, Widdel F (1993) Complete oxidation of toluene under strictly anoxic conditions by a new sulfate-reducing bacterium. Appl Environ Microbiol 59: 1444–1451. Rabus R, Wilkes H, Behrends A, Armstroff A, Fischer T, Pierik AJ, Widdel F (2001) Anaerobic initial reaction of n-alkanes: evidence for (1-methylpentyl)succinate as initial product and for involvement of an organic radical in the metabolism of n-hexane in a denitrifying bacterium. J Bacteriol 183: 1707–1715.

Energetic and Other Quantitative Aspects of Microbial Hydrocarbon Utilization Reardon KF, Mosteller DC, Bull Rogers JD (2000) Biodegradation kinetics of benzene, toluene, and phenol as single and mixed substrates for Pseudomonas putida F1. Biotechnol Bioeng 69:385–400. Russell JB (2007) The energy spilling reactions of bacteria and other organisms. J Mol Microbiol Biotechnol 13: 1–11. Schink B (1997) Energetics of syntrophic cooperation in methanogenic degradation. Microbiol Mol Biol Rev 61: 262–280. Schink B (2002) Anaerobic digestion: concepts, limits and perspectives. Water Sci Technol 45: 1–8. Shima S, Thauer RK (2005) Methyl-coenzyme M reductase and the anaerobic oxidation of methane in methanotrophic Archaea. Curr Opin Microbiol 8: 643–648. So CM, Young LY (1999) Isolation and characterization of a sulfate-reducing bacterium that anaerobically degrades alkanes. Appl Environ Microbiol 65: 2969–2976. Stouthamer AH (1988) Bioenergetics and yields with electron acceptors other than oxygen. In LE Erickson, DY-C Fung (eds.) Handbook of Anaerobic Fermentations. New York: Marcel Dekker, pp. 345–437. Stumm W, Morgan JJ (1981) Aquatic chemistry, 2nd edn. New York: John Wiley & Sons. Tan NC, van Doesburg W, Langenhoff AA, Stams AJ (2006) Benzene degradation coupled with chlorate reduction in a soil column study. Biodegradation 17: 113–119. Tempest DW, Neijssel OM (1984) The Status of YATP and maintenance energy as biologically interpretable phenomena. Ann Rev Microbiol 38: 459–486.

1

Thauer RK, Shima S (2008) Methane as a fuel for anaerobic microorganisms. Ann NY Acad Sci 1125: 158–170. Thauer RK, Jungermann K, Decker K (1977) Energy conservation in chemotrophic anaerobic bacteria. Bacteriol Rev 41: 100–180. Tissot BP, Welte DH (1984) Petroleum formation and occurrence. Berlin: Springer. van Dijken JP, Harder W (1975) Growth yields of microorganisms on methanol and methane. A theoretical study. Biotechnol Bioeng 17:15–30. Wagner F, Kleemann T, Zahn W (1969) Microbial transformations of hydrocarbons. II. Growth constants and cell composition of microbial cells derived from n-alkanes. Biotechnol Bioeng 11: 393–408. Widdel F, Musat F, Knittel K, Galushko A (2007) Anaerobic degradation of hydrocarbons with sulphate as electron acceptor. In LL Barton, WA Hamilton (eds.) Sulphate-Reducing Bacteria. Cambridge: Cambridge University Press, pp. 265–303. Wilhelm E, Battino R, Wilcock RJ (1977) Low-pressure solubility of gases in liquid water. Chem Rev 77: 219–262. Wodzinski RS, Johnson MJ (1968) Yields of bacterial cells from hydrocarbons. Appl Microbiol 16: 1886–1891. Yamamoto S, Alcauskas JB, Crozier TE (1976) Solubility of methane in distilled water and seawater. J Chem Eng Data 21: 78–80. Zengler K, Richnow HH, Rosello´-Mora R, Michaelis W, Widdel F (1999) Methane formation from longchain alkanes by anaerobic microorganisms. Nature 401: 266–269.

763

Part 2

Biochemistry of Aerobic Degradation

2 Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria T. J. Smith1 . Y. A. Trotsenko2 . J. C. Murrell3 1 Biomedical Research Centre, Sheffield Hallam University, Sheffield, UK 2 G.K.Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino, Moscow Region, Russia 3 Department of Biological Sciences, University of Warwick, Coventry, UK [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 768

2

Physiology and Biochemistry of Methanotrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 769

3

Methane Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 769

4

Methanol Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 771

5

Formaldehyde and Formate Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 771

6

Carbon Assimilation Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 772

7

Nitrogen Metabolism in Methanotrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 776

8

The Obligate Nature of Methanotrophs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 776

9

Methanotrophs and Biotechnology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 777

10 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 778

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_58, # Springer-Verlag Berlin Heidelberg, 2010

768

2

Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

Abstract: Methanotrophic bacteria grow aerobically using methane as a source of carbon and energy. They are widespread in the environment and play an important role in oxidizing methane in the environment, thereby mitigating the effects of global warming by this potent greenhouse gas. Methane monooxygenases (MMOs), which are the enzymes that catalyze the oxidation of methane, especially, the catalytically versatile soluble MMO, can cooxidize a wide range of hydrocarbons and chlorinated hydrocarbons, and have great potential as biocatalysts for bioremediation and biocatalysis. Methanotrophs can also be used to make single-cell protein from methane. Recent isolation of novel groups of thermophilic, acidophilic methanotrophs has revealed that these bacteria can even grow under extreme environmental conditions. The availability of genome sequences of several methanotrophs now opens up possibilities of postgenomic studies to investigate the regulation of methane oxidation in the laboratory and in the environment.

1

Introduction

Methane-oxidizing bacteria (methanotrophs) are a widely distributed group of aerobic microorganisms that use methane as their sole source of carbon and energy. They synthesize all of their cellular carbon-containing molecules from methane, and can also co-oxidize a wide range of hydrocarbons (Dalton, 2005). Anaerobic methane-oxidizing bacteria also exist but only the properties and metabolism of the aerobic methanotrophs will be described in this chapter. The majority of methanotrophs are Gram-negative bacteria that are classified as type I or type II methanotrophs according to whether they belong to the g- or a-subdivisions of the proteobacteria, respectively. They play a major role in oxidizing methane that is produced in the environment due to the activity of methanogenic archaea, and therefore play a major role in the global methane cycle by preventing the release of much of this methane to the atmosphere, thereby mitigating the effects of this potent greenhouse gas (Hanson and Hanson, 1996). Methanotrophs contain the remarkable enzyme methane monooxygenase (MMO). MMOs oxidize methane to methanol and catalyze a very large number of adventitious oxidation reactions that are the basis of the ability of methanotrophs to carry out processes in biocatalysis and bioremediation. Thus, they have found applications in production of single cell protein (SCP) from natural gas and bioremediation of trichloroethene (TCE)contaminated groundwater. Methanotrophs are ubiquitous in nature and have been isolated from many environments including soils, peatlands, rice paddies, sediments, freshwater and marine systems, acidic hot springs, mud pots, alkaline soda lakes, cold environments, and tissues of higher organisms (McDonald et al., 2008). Methanotrophs can broadly be divided into Type I and Type II methanotrophs (g- and a-proteobacteria, respectively). Genera of Type I methanotrophs include Methylomonas, Methylobacter, Methylococcus, Methylosphaera, Methylocaldum, Methylothermus, Methylohalobius, Methylomicrobium, Methylosarcina, and Methylosoma and Type II methanotrophs include Methylosinus, Methylocystis, Methylocella, and Methylocapsa. Recently two filamentous methanotrophs, Crenothrix polyspora and Clonothrix fusca as well as the first fully authenticated facultative methanotroph, Methylocella silvestris, which can grow on either methane or some multi-carbon compounds, have been described (Dedysh et al., 2005; Theisen and Murrell, 2005). Even more remarkable is the recent isolation of three thermoacidophilic methanotrophs which belong to the bacterial phylum Verrucomicrobia and thus are only distantly related to the proteobacterial methanotrophs. These three new isolates provisionally named ‘‘Methylokorus’’, ‘‘Acidimethylosilex,’’ and ‘‘Methyloacida’’, grow remarkably at pH 1.5

Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

2

and 65 C (reviewed in Semrau et al., 2008). The characteristics of these aerobic methanotrophs are summarized in > Table 1 and described in > Chapter 26, Vol. 3, Part 1.

2

Physiology and Biochemistry of Methanotrophs

MMO, the principal defining enzyme of methanotrophs, exists in two structurally and biochemically distinct forms, particulate (pMMO) and soluble (sMMO). pMMO is a coppercontaining enzyme that is associated with unusual intracellular membranes that take the form of vesicular disks in type I methanotrophs and paired peripheral layers in type II organisms. sMMO is a cytoplasmic di-iron center-containing enzyme complex (reviewed in Hakemian and Rosenzweig, 2007). Both forms of MMO, soluble (sMMO) and particulate (pMMO), can not only oxidize methane to methanol but also co-oxidize a range of hydrocarbons and chlorinated pollutants, and hence are responsible for much of the biotechnological potential of methanotrophs (Smith and Dalton, 2004). In methanotrophs that can express either form of MMO, expression is regulated by the availability of copper ions. The copper-containing pMMO is expressed during growth at high copper-to-biomass ratios and sMMO is expressed during growth under low copper-to-biomass ratios (Murrell et al., 2000). The biochemistry and molecular biology of methane monooxygenases is described > Chapter 17, Vol. 2, Part 4 and will only briefly be outlined below. Methanol, the initial oxidation product of methane, is oxidized to formaldehyde by a PQQ-dependent methanol dehydrogenase (MDH) (Anthony and Williams, 2003). Formaldehyde is an important branch-point in methylotrophic metabolism and multiple pathways for metabolism of formaldehyde are common in methanotrophs. The methanotrophs possess two pathways for fixation of formaldehyde, the serine and ribulose monophosphate (RuMP) cycles, which are active in type I and type II methanotrophs, respectively (Anthony, 1982; Trotsenko and Murrell, 2008). In addition, some methanotrophs can fix carbon dioxide into biomass via the ribulose-bis-phosphate (RuBP) cycle and its key enzyme RuBP carboxylase/oxygenase (Rubisco).

3

Methane Oxidation

sMMO is a three-component binuclear iron center monooxygenase that belongs to a large group of bacterial hydrocarbon oxygenases known as the soluble di-iron monooxygenases. sMMO, encoded by a six-gene operon mmoXYBZDC, has three components: (1) a 250-kDa hydroxylase with an (abg)2 structure in which the a-subunits (MmoX) contain the binuclear iron active center, where substrate oxygenation occurs; (2) a 39-kDa NAD(P)H-dependent reductase (MmoC); (3) a 16-kDa component (MmoB) known as protein B or the coupling protein that contains no prosthetic groups or metal ions. There are X-ray crystal structures for the hydroxylase component and NMR data on the structures of protein B and the flavin domain of the reductase. The catalytic cycle of sMMO has been extensively studied and excellent progress has been made toward understanding the mechanism of oxygen and hydrocarbon activation at the di-iron center (Baik et al., 2003). The crucial reaction intermediate compound Q accumulates when the reduced (FeII-FeII) hydroxylase is reacted with O2 in the presence of protein B. In compound Q, which is kinetically competent to oxidize methane and other substrates, the di-iron center is most likely in the diferryl (FeIV-FeIV) state. The site of substrate binding of sMMO is a hydrophobic cavity deeply buried in the a subunit of the hydroxylase. There is now a good expression system for sMMO and structural and

769

Verrucomicrobia

Verrucomicrobia

Acidimethylosilex

Methyloacida

ND

pMMO

pMMO

pMMO

pMMO

pMMO

pMMO +/– sMMO

pMMO

pMMO

pMMO

pMMO + sMMO

pMMO

pMMO

pMMO +/– sMMO

pMMO +/– sMMO

b

ICM, intracellular membrane PLFA, phospholipid fatty acid c ND, not determined d NA, not applicable because ICMs are very limited in this genus

a

Verrucomicrobia

a Proteobacteria

Methylosinus

g Proteobacteria

a Proteobacteria

Methylocystis

Methylokorus

g Proteobacteria

Methylohalobius

Clonothrix

g Proteobacteria

Methylothermus

g Proteobacteria

g Proteobacteria

Methylocaldum

Crenothrix

sMMO

g Proteobacteria

Methylococcus

a Proteobacteria

g Proteobacteria

Methylosphaera

a Proteobacteria

g Proteobacteria

Methylosarcina

Methylocella

g Proteobacteria

Methylomonas

Methylocapsa

pMMO + sMMO

g Proteobacteria

Methylomicrobium

pMMO

g Proteobacteria

Methylosoma

pMMO

g Proteobacteria

MMO type

Methylobacter

Phylogeny

ND

RuMP?

Serine

ND

ND

Serine

Serine

Serine

Serine

RuMP

RuMP

RuMP/Serine

RuMP/Serine

RuMP

RuMP

RuMP

RuMP

not known

RuMP

C1 assimilation

NA

type IV?

type IV?

type IV?

type I

type I

type III

d

type II

type II

type I

type I

type I

type I

NDc

type I

type I

type I

type I

type I

ICM typea

Yes

No

No

ND

ND

Yes

Yes

Yes

Yes

No

No

No

Yes

Yes

No

some

No

Yes

No

N2 fixation

ND

ND

ND

ND

ND

63.1

60–61

63–67

62–67

58.7

62.5

57

59–66

43–46

54

51–59

49–60

49.9

49–54

G + C (mol %)

ND

C18:0

ND

ND

ND

18:1

18:1

18:1

18:1

18:1

18:1/16:0

16:1

16:1

16:1

16:1

16:1

16:1

16:1

16:1

Major PLFAb

Thermoacidophilic

Thermoacidiophilic

Thermoacidophilic

Not extreme

Not extreme

Acidophilic

Acidophilic

Not extreme

Some acidophilic

Halophilic

Thermophilic

Thermophilic

Thermophilic

Psychrophilic

Not extreme

Some psychrophilic

Halotolerant; alkaliphilic

Not extreme

Some psychrophilic

Trophic niche

2

Genus

. Table 1 Classification of genera of aerobic methanotrophs

770 Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

2

site-directed mutagenesis studies have indicated that a gating residue, Leucine 110, in the a subunit of the hydroxylase is important in determining how substrates enter and are presented at the active site (Borodina et al., 2007). pMMO is a copper-containing, membrane-associated enzyme that consists of three polypeptides, of 49, 27, and 22 kDa, encoded by the genes pmoB, A, and C, respectively. The X-ray crystal structure of pMMO shows that the enzyme has an (abg)3 stoichiometry (reviewed in Lieberman and Rosenzweig, 2005). The enzyme complex contains copper ions and also a di-iron center. A powerful copper chelator, methanobactin, is used to sequester copper ions for activity and/or structural integrity of pMMO (reviewed in Balasubramanian and Rosenzweig, 2008). The best characterized methanotrophs, Methylococcus capsulatus (Bath) (type I) and Methylosinus trichosporium OB3b (type II), can produce either form of MMO. During growth under high copper-to-biomass ratios, pMMO is expressed whereas the soluble form of the enzyme is expressed when the copper-to-biomass ratio is low (Murrell et al., 2000). Methanotrophs that possess only pMMO, such as Methylomonas methanica and Methylomicrobium album BG8 have been known for many years. Recently the facultative type II methanotrophs Methylocella silvestris has been shown to possess sMMO but not pMMO. The properties of MMOs have attracted interest for biotechnological applications and these are described in Section 9.

4

Methanol Oxidation

Methanol produced from the oxidation of methane is further oxidized by a periplasmic enzyme methanol dehydrogenase (MDH) which has pyrroloquinoline quinone (PQQ) as a prosthetic group (Anthony and Williams, 2003). MDH is a heterotetramer a2b2 consisting of two large (67 kDa) and two small (8.5 kDa) subunits. Oxidation of methanol is coupled with reduction of the prosthetic group (PQQH2) into the corresponding quinol (PQQH2) followed by two-step transfer of electrons to the acceptor which is an inducible cytochrome c551 (cL) and further via the cytochromes c550 (cn), and c552 to the terminal oxidase. The biochemistry and molecular biology of methanol oxidation has been mainly studied in the Gram-negative methanol-utilizing bacterium Methylobacterium extorquens (Chistoserdova et al., 2003) but it is likely that the same mechanisms operate in methanotrophs. The components of the methanol oxidation pathway are encoded by at least 25 genes. mxaJ, mxaR and mxaS, mxaD, mxaE, and mxaH are required for the formation of active MDH, while mxaG encodes cytochrome c551 and the products of genes mxaACKLD are involved in insertion of PQQ and Ca2+into MDH. The polypeptide encoded by mxaB regulates transcription of the MDH genes. Two gene clusters, pqqABCDE and pqqFG, are involved in biosynthesis of PQQ. pqqA encodes the proposed precursor of PQQ, containing tyrosine and glutamate. mxbDM and mxaQE, are required for transcriptional regulation of the methanol oxidation system.

5

Formaldehyde and Formate Oxidation

Formaldehyde is the key intermediate in the linear pathway of methane oxidation to CO2. A major portion of the reducing equivalents required for methane oxygenation is formed during formaldehyde oxidation via formate to CO2. Methanotrophs have several enzymes

771

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Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

involved in formaldehyde oxidation. In Mc. capsulatus (Bath) expressing pMMO, formaldehyde is mainly oxidized by a particulate cytochrome-linked formaldehyde dehydrogenase (FADH). Again, the majority of recent work done on formaldehyde and formate oxidation has been with methylotrophs which use pterin cofactors, H4F/THF and H4MPT, for activation of formaldehyde oxidation and probably the same mechanisms operate in methanotrophs. In Type II methanotrophs, the major role of the THF-pathway enzymes is maintenance of a high concentration of N5,N10-methylene-THF, which is the primary acceptor of formaldehyde in the serine pathway of C1 assimilation. Because of the reversibility of the reactions catalyzed, these enzymes can be regulated by the requirements of the cell either into the oxidation or assimilation of formaldehyde (Vorholt, 2002). The functions of the THF-pathway enzymes have not been elucidated in Type I methanotrophs (> Figs. 1 and 2). The reactions of H4MPT-dependent oxidation of formaldehyde were first found in methanogenic and sulfate reducing Archaea and were considered to be a specific characteristic only of these strict anaerobes, till their discovery in the aerobic facultative methylotroph Methylobacterium extorquens AM1. The highest activities of the H4MPT pathway enzymes, methenyl-H4MPT cyclohydrolase and NAD(P)-dependent methenyl-H4MPT dehydrogenase, are found in Mc. capsulatus (Bath) and other methanotrophs of the a- and g-proteobacteria. MethenylH4MPT-cyclohydrolase (Mch) catalyzes conversion of methenyl-H4MPT into N10-formyl-H4MPT. Formate oxidation is a final step in the methane oxidation pathway. In all extant methanotrophs, an NAD+ -dependent formate dehydrogenase (FDH) is present (reviewed in Trotsenko and Murrell, 2008). The enzyme from Ms. trichosporium OB3b consists of two types of polypeptides (54 and 106 kDa) and functions in vitro as an electron donor for sMMO or nitrogenase (with the additional participation of ferredoxin-NAD+ reductase and ferredoxin). In cell extracts of many methanotrophs, a highly active phenazine methosulfate-linked FDH associated with membranes has been found. In the genome of Mc. capsulatus (Bath) there are open reading frames (ORFs) encoding FDH-like proteins and their role warrants further study.

6

Carbon Assimilation Pathways

The pioneering studies of Prof J.R.Quayle and colleagues (e.g., see Strom et al., 1974) revealed that methanotrophs use two major pathways for primary C1 assimilation, the RuMP and serine cycles. In both of these cycles, phosphotrioses are synthesized from formaldehyde which is the key intermediate in carbon metabolism (Anthony, 1982). In the first part of the RuMP cycle, formaldehyde is fixed with ribulose-5-phosphate to form (D-arabino)-3-hexulose-6-phosphate in a reaction catalyzed by 3-hexulosephosphate synthase (HPS). This very unstable product is rapidly isomerized to fructose-6-phosphate by phosphohexuloisomerase (PHI). These two specific enzymes catalyze the formation of both C-C bonds and phosphohexoses in Type I methanotrophs. In Mc. capsulatus (Bath), the purified HPS is a homohexameric (310 kDa) membrane-bound enzyme with an unusually large subunit size (49 kDa). Based on its gene sequence, the HPS molecular mass appears to correspond to the product of the hps-phi-fused gene. By comparison with HPS, the characteristics of PHI are poorly documented in methanotrophs. In the second part of the RuMP cycle, the phosphohexoses are split to (phospho)trioses. In methanotrophs this cleavage occurs by two simultaneous pathways, via the Entner-Doudoroff and Embden-Meyerhof-Parnas variants. In the first variant, fructose-6-phosphate is converted via glucose-6-phosphate

Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

. Figure 1 Pathways of carbon metabolism in Type I methanotrophs.

2

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774

2

Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

. Figure 2 Pathways of carbon metabolism in Type II methanotrophs.

and 6-phosphogluconate into 2-keto-3-deoxy-6-phosphogluconate (KDPG), which is subsequently cleaved by KDPG-aldolase to pyruvate and glyceraldehyde-3-phosphate (GAP). Alternatively, in the second variant fructose-6-phosphate is phosphorylated by ATP into fructose1,6-bisphosphate, followed by aldolase cleavage to GAP and dihydroacetonephosphate, the latter being isomerized to GAP. Nonetheless, the glycolytic cleavage of phosphohexoses into phosphotrioses has not been considered as a physiologically significant pathway in methano-

Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

2

trophs because of the low or zero activity of ATP-dependent 6-phosphofructokinase (Strom et al., 1974). However, the discovery of very active pyrophosphate-dependent 6-phosphofructokinase (PPi-PFK), together with the high intracellular levels of PPi (up to 20 mM), have established that PPi, but not ATP, is the phosphoryl donor in this. Furthermore, the purified PFK of Methylomonas methanica 12, a homodimer of 2  45 kDa, is rather similar in kinetic and regulatory properties to the analogous enzymes from microbes rather than plants. The recent identification and cloning of the pfk gene revealed its distant similarity (16.5% identical amino sequences) to that of Mc. capsulatus (Bath) and the facultative methylotroph Amycolatopsis methanolica (23.2%). This finding suggests a different origin and metabolic role for the PPi-PFKs among these bacteria. The high degree of divergence of the pfk gene in methanotrophs may be determined by the characteristics of primary and central metabolism among these bacteria, as well as with differences in the function and activity of the PPi-PFK. Significantly, PPi-PFK having a crucial position at a metabolic cross-roads catalyzes the easily reversible interconversion of fructose-6-phosphate into fructose-1,6-bisphosphate and participates in distribution of the carbon flux between the glycolytic and KDPG segments (branches) of the RuMP cycle. In fact, this metabolic loop serves to balance the levels of GAP and (phosphoenol) pyruvate in the cell (Trotsenko and Murrell, 2008). In the third part of the RuMP cycle, the primary acceptor of formaldehyde, i.e., ribulose-5P is regenerated from glyceraldehyde-3P and fructose-6P in a series of transaldolase/transketolase reactions analogous to photo- and chemotrophic bacteria. Type II methanotrophs employ the serine cycle for C1-assimilation. In the first part of the serine cycle, formaldehyde (after condensation with THF and formation of N5,N10-methylene THF) reacts with glycine to produce serine by the action of the appropriate serinehydroxytransmethylase (SHTM). The amino group of serine is then transferred by a specific serine-glyoxylate aminotransferase (SGAT) to glyoxylate thus forming glycine and hydroxypyruvate (by hydroxypyruvate reductase, HPR) which is phosphorylated by ATP-glycerate kinase to 2-phosphoglycerate, followed by isomerization to PEP and its subsequent carboxylation to oxaloacetate. The reduction of oxaloacetate, catalyzed by malate dehydrogenase, forms malate which is then converted to malyl-CoA by malate thiokinase. Finally, malyl-CoA lyase forms glyoxylate and acetyl-CoA, the latter being a primary product of the serine cycle. Consequently, SHTM, SGAT, HPR, and malyl-CoA lyase are the key indicative enzymes of the serine cycle. In the second part of the serine cycle, acetyl-CoA is oxidized to glyoxylate which is further (trans)aminated to glycine, so that the primary acceptor of formaldehyde is regenerated. Interestingly, the obligate methanotrophs and serine pathway methylotrophs ‘‘lacked’’ isocitrate lyase (icl- variant). As shown recently, glyoxylate can be regenerated via the formation of acetoacetyl-CoA and hydroxybutyryl-CoA, the known intermediates of the poly-bhydroxybutyrate biosynthesis pathway, and also crotonyl-CoA and butyryl-CoA, the intermediates of fatty acid biosynthesis. However, at present it is not clear whether this rather complicated pathway proposed for M. extorquens AM1 or other variants of this pathway (via citramalate or methylmalate) operates in Type II methanotrophs (Meister et al., 2005). Methanotrophs of the genera Methylococcus and Methylocaldum assimilate formaldehyde mainly by the RuMP pathway, although they also possess less active enzymes for the serine pathway and the Calvin-Benson-Basham (CBB) cycle, i.e., phosphoribulokinase and ribulosebisphosphate carboxylase/oxygenase (Rubisco). The role of these enzyme activities is still unclear. In Type I methanotrophs, 5–15% of the carbon in cell biomass is derived from CO2 while in Type II methanotrophs it is up to 50%. Moreover, in both Type I and Type II methanotrophs, PEP carboxylase is responsible for anaplerotic CO2 fixation (Shishkina and Trotsenko,

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Physiology and Biochemistry of the Aerobic Methane Oxidizing Bacteria

1982). In the Type I methanotrophs Methylococcus the CBB cycle enzymes are also involved in CO2 fixation. Rubisco of Mc. capsulatus (Bath) has an a6b6 structure which differs from the typical hexadecameric structure (a8b8) of the Form I Rubisco formed in Proteobacteria and also in cyanobacteria and higher plants. The Rubisco genes encoding the large subunit (cbbL), small subunit (cbbS) and putative regulatory gene (cbbQ) are located on one cluster in Methylococcus (Ward et al., 2004; reviewed in Kelly et al., 2005).

7

Nitrogen Metabolism in Methanotrophs

The ability to fix N2 by Type II methanotrophs and Mc. capsulatus (Bath) has been known for some time. More recently, the potential for other Type I methanotrophs such as Methylomonas and Methylobacter to fix N2 has also been determined by screening for the gene nifH which encodes the Fe-containing protein of nitrogenase. The structural genes for nitrogenase of Mc. capsulatus (Bath) (nifH, nifD and nifK) are contiguous as they are in other diazotrophs and analysis of the genome of Mc. capsulatus has revealed the presence of other nif genes involved in synthesis and maturation of the nitrogenase iron-molybdenum cofactor. Type I methanotrophs assimilated NH4+ mainly by reductive amination of pyruvate and/ or a-ketoglutarate, whereas Type II methanotrophs used the glutamate cycle, i.e., glutamine synthetase (GS) and the glutamine-oxoglutarate amidotransferase (GOGAT) system. The GS purified from Mc. capsulatus (Bath) is regulated by (de)adenylylation mechanisms. At concentrations of > 0.5 mM NH4 in the medium, GS exists in the non-active adenylylated form. The structural gene for GS from Mc. capsulatus (Bath) has been cloned and sequenced. Its nucleotide sequence has 59% similarity with glnA gene of Anabaena sp. 7120. Regulation of glnA in this methanotrophs is analogous to that of enterobacteria and occurs via the Ntr system. In Mc. capsulatus (Bath) and Type I methanotrophs grown on medium containing ammonia, the reductive amination of pyruvate (via alanine dehydrogenase) and/or a-ketoglutarate (via glutamate dehydrogenase) occurs under high ammonia growth conditions. In contrast, when grown under N2-fixing conditions, i.e., under ammonium limitation ( Chapter 23, Vol. 2, Part 5. However, some bacterial species have been characterized in the last few years that are highly specialized in degrading hydrocarbons. They are called hydrocarbonoclastic bacteria and play a key role in the removal of hydrocarbons from polluted environments (Harayama et al., 2004; Head et al., 2006; Yakimov et al., 2007). Of particular importance is Alcanivorax borkumensis, a marine bacterium that can assimilate linear and branched alkanes, but which is unable to use aromatic hydrocarbons, sugars, amino acids, fatty acids and most other common substrates as the carbon source (Schneiker et al., 2006; Yakimov et al., 1998). Alcanivorax sp. are present in nonpolluted sea waters in low numbers, probably living at the expense of the alkanes that are continuously produced by algae and other sea organisms and that are present at low but constant concentrations. Alcanivorax strains become predominant after a spill of crude oil and are believed to play an important role in natural bioremediation of oil spills worldwide

Enzymes for Aerobic Degradation of Alkanes

3

(Hara et al., 2003; Harayama et al., 2004; Kasai et al., 2002; McKew et al., 2007a, b; Yakimov et al., 2007). Hydrocarbonoclastic alkane-degrading bacteria of the genera Thalassolituus (Yakimov et al., 2004), Oleiphilus (Golyshin et al., 2002) and Oleispira (Yakimov et al., 2003) also play an important role in the biodegradation of oil spills in several environments (Coulon et al., 2007; McKew et al., 2007a, b). Alkanes can be metabolized aerobically or anaerobically. This chapter deals only with aerobic degradation, since anaerobic degradative pathways are covered in a separate chapter in this book. The pathways and enzymology for the degradation of alkanes have been reviewed extensively before (Ashraf et al., 1994; Coon, 2005; Rehm and Reiff, 1981; van Beilen and Funhoff, 2007; van Hamme et al., 2001; Watkinson and Morgan, 1990; Wentzel et al., 2007). This chapter will emphasize the most recent developments on alkane metabolism. Regulation of the expression of the genes involved in alkane degradation is treated on a separate chapter in this handbook. The degradation of methane, which is a special case that is oxidized by a very specialized enzyme, is also covered on a separate chapter.

2

Uptake of n-Alkanes

The water solubility of n-alkanes decreases exponentially as their molecular weight increases (Eastcott et al., 1988; see > Table 1). For alkanes having more than nine carbon atoms, solubility is negligible from the point of view of their uptake by microorganisms. This poses a problem for their biodegradation. The precise way in which alkanes enter the cell is unclear, but the mechanism probably differs depending on the bacterial species considered, the molecular weight of the alkane and the physico-chemical characteristics of the environment (Wentzel et al., 2007). The direct uptake of the alkane molecules from the water phase can only be considered as a relevant possibility for low molecular weight alkanes, which are still sufficiently soluble to assure a proper mass transfer to the cell. For medium- and long-chain n-alkanes, microorganisms may gain access to these compounds either by adhering to hydrocarbon droplets (which is facilitated by a hydrophobic cell surface) or by a surfactantfacilitated access. Most bacteria able to degrade n-alkanes produce and secrete surfactants of diverse chemical nature that allow emulsification of the hydrocarbons (Hommel, 1990; Ron and Rosenberg,

. Table 1 Water solubility of representative n-alkanes (at 25 C) Solubility (mol L 1)

n-Alkane

Carbon atoms

Molecular weight

Propane

3

44.1

510

Hexane

6

86.2

1.410 6

Nonane

3 4

9

128.3

10

Dodecane

12

170.3

210

8

Hexadecane

16

226.4

210

10

12

Eicosane

20

282.6

10

Hexacosane

26

366.7

410

Data obtained from Eastcott et al. (1988)

16

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784

3

Enzymes for Aerobic Degradation of Alkanes

2002). Biosurfactants are believed to increase the surface area that hydrophobic compounds can expose to the water phase, thereby facilitating the access of microorganisms to the oil phase (Ron and Rosenberg, 2002). In liquid cultures, surfactants have been reported to increase the uptake and assimilation of alkanes such as hexadecane (Beal and Betts, 2000; Noordman and Janssen, 2002). However, in soils and other situations the usefulness of surfactants for the uptake of alkanes is less evident (Holden et al., 2002). It should be noted that efficient emulsification requires the production of relatively large amounts of the surfactant, which in turn requires high population densities of the surfactant-producing microorganism. This suggests that the role of surfactants at low cell densities could be different from emulsification. P. aeruginosa produces a rhamnolipid surfactant that stimulates the uptake of hexadecane through a process that requires energy (Beal and Betts, 2000; Noordman and Janssen, 2002). Uptake of the alkanes may also occur through their passive diffusion into the cell membrane. Surfactants produced by microorganisms probably serve other biological functions in addition to emulsifying hydrocarbons to improve their uptake (Ron and Rosenberg, 2001). For example, surfactants facilitate adhesion to and detachment from surfaces of from biofilms (Boles et al., 2005; Neu, 1996), as well as cell motility on solid surfaces (Caiazza et al., 2005; Kohler et al., 2000). In the case of alkane degrading bacteria that also behave as opportunistic pathogens, like Pseudomonas aeruginosa, these properties of biosurfactants can facilitate certain infections, so that they can also be considered as a virulence factor (Zulianello et al., 2006). In other words, bacteria can use surfactants for several purposes in different environmental conditions, the uptake of hydrocarbons being just one of the processes where the properties of surfactants can be useful.

3

Pathways for Degradation of n-Alkanes

In most cases described, aerobic degradation of n-alkanes starts by the oxidation of a terminal methyl group to render a primary alcohol, which is further oxidized to the corresponding aldehyde, and finally converted into a fatty acid (see > Fig. 1). Fatty acids are conjugated to CoA and further processed by b-oxidation to generate acetyl-CoA (Ashraf et al., 1994; Rehm and Reiff, 1981; van Hamme et al., 2003; Watkinson and Morgan, 1990; Wentzel et al., 2007). Subterminal oxidation has been reported as well in some microorganisms (> Fig. 1; Britton, 1984; Kotani et al., 2003, 2006; Whyte et al., 1998). While oxidation of fatty alcohols and fatty acids is common among microorganisms, activation of the alkane molecule requires an enzyme system that is much less widespread.

4

Hydroxylation of n-Alkanes

In bacteria, the initial terminal hydroxylation of n-alkanes can be carried out by enzymes belonging to different families (> Table 2; van Beilen and Funhoff, 2007; van Beilen et al., 2003). Microorganisms degrading short-chain-length alkanes (C2–C4, where the subindex indicates the number of carbon atoms of the alkane molecule) have enzymes related to methane monooxygenases. Strains degrading medium-chain-length alkanes (C5–C11), or long-chain-length alkanes (>C12), frequently contain integral membrane non-heme iron monooxygenases related to the well-characterized Pseudomonas putida GPo1 AlkB alkane

3

Enzymes for Aerobic Degradation of Alkanes

. Figure 1 Most frequent pathways for the degradation of n-alkanes by terminal and subterminal oxidation.

. Table 2 Enzyme classes oxidizing alkanes Enzyme class

Characteristics

Substrate length

Host

PRM, propane monooxygenase

Non-heme iron monooxygenase similar to sMMO

C3

Bacteria

sBMO, butane monooxygenase

Non-heme iron monooxygenase similar to sMMO

C2–C9

Bacteria

pBMO, butane monooxygenase

Copper-containing monooxygenase similar to pMMO

C2–C9

Bacteria

CYP153

Soluble cytochrome P450

C5–C12

Bacteria

CYP52

Membrane-bound cytochrome P450

C10–C16

Yeasts

AlkB-related

Non-heme iron monooxygenase

C3–C13 or C10–C20

Bacteria

AlmA

Flavin-binding monooxygenase

C20–C36

Bacteria

LadA

Thermophilic flavin-dependent monooxygenase

C10–C30

Bacteria

Dioxygenase

Copper flavin-dependent dioxygenase

C10–C30

Bacteria

The substrate range is approximate; upper and lower limits may vary in different strains. See text for details

hydroxylase. However, some strains contain alkane hydroxylating enzymes that belong to a family of soluble cytochrome P-450s and that are active against C5–C11 alkanes. Finally, several strains assimilating alkanes of more than 18 carbon atoms contain alkane hydroxylases that seem to be unrelated to the former ones and that only recently have started to be characterized.

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Enzymes for Aerobic Degradation of Alkanes

Several yeasts can assimilate alkanes as well. In those cases studied, the enzymes involved in the initial oxidation of the alkane molecule belong to the family of microsomal cytochrome P450 (Iida et al., 2000; Ohkuma et al., 1998; Zimmer et al., 1996). The role of yeasts in the biodegradation of n-alkanes in oil-contaminated sites may be more significant than previously considered, at least in some environments (Schmitz et al., 2000).

4.1

Alkane Hydroxylases Related to Methane Monooxygenase

Several bacterial strains can grow on C2–C4 gaseous alkanes, but not on methane (Ashraf et al., 1994). The enzymes that initially oxidize these alkanes are related to methane monooxygenases (Hamamura et al., 1999). There are two different forms of methane monooxygenases. All methanotrophs produce a membrane-bound particulate form of methane monooxygenase (pMMO) which oxidizes a narrow range of alkanes, while some methanotrophs additionally produce as well a soluble form of methane monooxygenase (sMMO) that is active on a broader range of substrates and oxidizes C1–C7 alkanes to the corresponding alcohols (Green and Dalton, 1989). Pseudomonas butanovora, which was later proposed to be a close relative to Thauera sp. on the basis of 16S DNA sequence (Anzai et al., 2000), can grow on C2–C4 alkanes by a pathway that sequentially oxidizes the terminal methyl group of the hydrocarbon (Arp, 1999). The first enzyme of the pathway, termed butane monooxygenase (BMO), is a non-heme iron monooxygenase similar to the sMMO and can hydroxylate C2–C9 alkanes (Sluis et al., 2002). The enzyme is composed of three components: a dinuclear ironcontaining monooxygenase (BMOH) that in turn contains three different polypeptides, an NADH-oxidoreductase (BMOR), and a small regulatory protein (BMOB) that probably acts as an effector and that may be partly dispensable (Dubbels et al., 2007). The proper assembly of BMO has been proposed to require the assistance of a chaperonin-like protein, BmoG (Kurth et al., 2008). Gordonia sp. TY-5, which can grow on propane as the carbon source, contains a propane monooxygenase that shows sequence similarity to sMMO, but has a very narrow substrate range; it can only oxidize propane and does so at the sub-terminal position, generating 2-propanol (Kotani et al., 2003). This secondary alcohol is then oxidized to acetone, which is further transformed into methylacetate and, finally, into acetic acid and methanol (Kotani et al., 2007). The genes encoding the Gordonia sp. TY-5 propane monooxygenase have been found as well in two propane-utilizing species, Mycobacterium sp. TY-6 and Pseudonocardia sp. TY-7 (Kotani et al., 2006). In Mycobacterium sp. TY-6, propane is oxidized at the terminal position. In Pseudonocardia sp. TY-7, however, both terminal and sub-terminal oxidation was observed. The butane monooxygenases of two other strains, Mycobacterium vaccae JOB5 and Nocardioides CF8, have been analyzed from a physiological point of view. In the absence of DNA sequence data, it seems that M. vaccae JOB5 butane monooxygenase shows properties similar to sMMO (Hamamura et al., 1999), while that of Nocardioides CF8 is a coppercontaining enzyme similar to pMMO (Hamamura and Arp, 2000; Hamamura et al., 1999).

4.2

The AlkB Family of Alkane Hydroxylases

The most extensively characterized alkane degradation pathway is that encoded on the OCT plasmid of P. putida GPo1, formerly identified as Pseudomonas oleovorans GPo1. It was

Enzymes for Aerobic Degradation of Alkanes

3

originally characterized by Coon and coworkers (Baptist et al., 1963) and has later become a model system (van Beilen et al., 1994, 2001). The first enzyme of this pathway is an integralmembrane non-heme diiron monooxygenase, named AlkB, that hydroxylates alkanes at the terminal position. AlkB requires two soluble electron transfer proteins named rubredoxin (AlkG) and rubredoxin reductase (AlkT). Rubredoxin reductase, via its cofactor FAD, transfers electrons from NADH to the rubredoxin, which in turn transfers the electrons to AlkB (see > Fig. 2). The biochemical properties of AlkB have been analyzed in detail. Although the crystal structure is not available, several approaches allowed to deduce that it has six transmembrane segments and a catalytic site that faces the cytoplasm. The active site includes four histidine-containing sequence motives that are conserved in other hydrocarbon monooxygenases and chelate two iron atoms (> Fig. 3; Shanklin et al., 1994; van Beilen et al., 1992b). The diiron cluster allows the oxygen-dependent activation of the alkane molecule through a substrate radical intermediate (Austin et al., 2000; Bertrand et al., 2005; Shanklin et al., 1997). One of the oxygen atoms of O2 is transferred to the terminal methyl group of the alkane, rendering an alcohol, while the other oxygen atom is reduced to H2O by electrons transferred by the rubredoxin. Oxidation is regio-and stereospecific (van Beilen et al., 1996). The P. putida GPo1 AlkB alkane hydroxylase can oxidize propane, n-butane (Johnson and Hyman, 2006), as well as C5–C13 alkanes (van Beilen et al., 2005b). All these alkanes can also support growth. Methane, ethane, or alkanes longer than C13, are not oxidized. A mutagenesis approach allowed identifying a residue, Trp55, which appears to limit the size of the alkane molecule that AlkB can oxidize, since when replaced by Ser or Cys the substrate range increased to include C14 and C16 alkanes (van Beilen et al., 2005b). It was proposed that the AlkB active site might be a deep hydrophobic pocket formed by the proper alignment of the six transmembrane helices, and that the alkane molecule should slide into it until the terminal methyl group is correctly positioned relative to the His residues that chelate the iron atoms (> Fig. 3). The estimated distance between the residue Trp55 and the His residues is similar to the length of a linear C13 molecule. This suggests that the bulky side chain of Trp55 would

. Figure 2 Oxidation of n-alkanes by alkane hydroxylases belonging to the AlkB family (left) or to the bacterial cytochrome P450 family (right). AH, membrane bound alkane hydroxylase; Rub, rubredoxin; RubR, rubredoxin reductase; Cyp P450, soluble cytochrome P450; Fdx, ferredoxin; FdxR, ferredoxin reductase. The gray bar represents the cytoplasmic membrane; the phospholipid layer facing the cytoplasm is marked as ‘‘In.’’

787

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Enzymes for Aerobic Degradation of Alkanes

. Figure 3 Model proposed for the structure of the P. putida GPo1 membrane-bound AlkB alkane hydroxylase. The gray bar represents the cytoplasmic membrane. The four histidine clusters (H) believed to bind the two iron atoms (Fe) at the catalytic site are indicated, as well as the proposed position of residue Trp55, which would extend its bulky side group to the hydrophobic pocket in which the alkane molecule is believed to fit in place. Adapted from van Beilen et al. (2005b) and Rojo (2005).

protrude into the hydrophobic pocket, impeding alkanes longer than C13 to enter deeper into the pocket, thereby impairing the proper alignment of the terminal methyl group with the catalytic site. The presence at position 55 of amino acids with a less bulky side chain would allow larger alkanes to fit in place into the hydrophobic pocket. More than sixty AlkB homologs are known to date (Marı´n et al., 2001, 2003; Smits et al., 1999, 2002, 2003; van Beilen et al., 2002b, 2004). They have been found in both Gram-positive and Gram-negative microorganisms and show a high sequence diversity (van Beilen et al., 2003). Interestingly, only a few of these AlkB enzymes oxidize C5–C13 alkanes, as does P. putida GPo1 AlkB, whereas most members of this family prefer alkanes larger than C10. The rubredoxin that transfers electrons to the AlkB active site is a small redox-active ironsulfur protein. The AlkG rubredoxin of P. putida GPo1 is unusual in that it contains two rubredoxin domains, AlkG1 and AlkG2, connected by a linker, while rubredoxins from other microorganisms have only one of these domains. Several rubredoxins present in Grampositive and Gram-negative alkane-degrading bacteria were cloned and analyzed in complementation assays for their ability to substitute for P. putida GPo1 AlkG. Interestingly, they clustered in two groups. AlkG1-type rubredoxins could not transfer electrons to the alkane hydroxylase, while AlkG2-type enzymes were able to do so and could substitute for GPo1 AlkG (van Beilen et al., 2002a). AlkG1-type rubredoxins probably have other as yet unknown roles. In fact, rubredoxin-rubredoxin reductase systems are present in many other organisms that are unable to degrade alkanes, where they serve other functions. For example, they play an important role in oxidative stress responses in anaerobic microorganisms by transferring reducing equivalents from NADH to superoxide reductases, or to rubredoxin:oxygen oxidoreductases, thereby reducing oxygen or reactive oxygen species (Frazao et al., 2000).

Enzymes for Aerobic Degradation of Alkanes

3

The structure of the rubredoxin-rubredoxin reductase complex, which has been solved in the case of Pseudomonas aeruginosa, seems to be highly optimized for rapid transport of reducing equivalents to the final receptor (Hagelueken et al., 2007).

4.3

Cytochrome P450 Alkane Hydroxylases

Cytochromes P450 are heme-thiolate proteins that catalyze the oxygenation of a large number of compounds. They are ubiquitous among all kingdoms of life and can be grouped in more than 100 families on the basis of sequence similarity. Almost all eukaryotic P450s are membrane-bound enzymes while most prokaryotic P450s are soluble. Several bacterial strains that degrade C5–C10 alkanes contain alkane hydroxylases that belong to a distinct family of bacterial soluble cytochrome P450 monooxygenases. The first member characterized was CYP153A1 from Acinetobacter sp. EB104 (Maier et al., 2001), but similar enzymes have been found in diverse strains of mycobacteria, rhodococci and proteobacteria (Sekine et al., 2006; van Beilen et al., 2005a, 2006). These cytochromes P450 require the presence of a ferredoxin and of a ferredoxin reductase that transfer electrons from NAD(P)H to the cytochrome (> Fig. 2). Complementation assays showed that many of these cytochrome P450 proteins can functionally substitute for P. putida GPo1 AlkB, showing that they are true alkane hydroxylases (van Beilen et al., 2006). The cytochrome P450 from Mycobacterium sp. HXN-1500 was purified and shown to hydroxylate C6–C11 alkanes to 1-alkanols with high affinity and regioselectivity (Funhoff et al., 2006). As stated above, several yeasts can assimilate alkanes. In those cases studied, the enzymes involved in the initial oxidation of the alkane molecule are membrane-bound cytochrome P450s of the CYP52 family (Iida et al., 2000; Ohkuma et al., 1998; Zimmer et al., 1996). They receive electrons from NADPH via FAD- and FMN-containing reductases.

4.4

Alkane Hydroxylases for Long-Chain n-Alkanes

Several bacterial strains have been reported to assimilate alkanes larger than C20 (for a compilation see Wentzel et al., 2007). The enzymes responsible for the oxidation of such alkanes, which are solid at room temperature, are still poorly characterized. In Acinetobacter sp. M1, which can grow on C13–C44 alkanes, several alkane oxidizing enzymes have been detected. Two of them, named AlkMa and AlkMb, are related to P. putida GPo1 AlkB and are membrane bound (Tani et al., 2001). A third enzyme has been reported that is soluble, requires Cu2+, does not receive electrons from NADH and is therefore clearly unrelated to the AlkB family of hydroxylases (Maeng et al., 1996). It has been proposed to be a dioxygenase that oxidizes C10–C30 alkanes generating n-alkyl hydroperoxides that render the corresponding aldehyde. A different Acinetobacter strain, named DSM 17874, also contains at least three alkane oxidizing enzymes. Two of them are AlkB paralogs similar to the AlkMa and AlkMb enzymes described above, and oxidize C10–C20 alkanes (Throne-Holst et al., 2006). A third enzyme has been reported that oxidizes C20 to >C32 alkanes. Its gene, designated almA, has been identified and codes for a flavin-binding monooxygenase (Throne-Holst et al., 2007). Genes homologous to almA were identified in several other long-chain n-alkane degrading strains, including Acinetobacter M1. Most notably, two genes similar to almA were also detected in the genome of A. borkumensis SK2.

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A different long-chain alkane hydroxylase has been characterized in the thermophilic bacterium Geobacillus thermodenitrificans NG80-2 (Feng et al., 2007). It is termed LadA and oxidizes C15–C36 alkanes, generating the corresponding primary alcohols. Its crystal structure has been solved, revealing that it belongs to the bacterial luciferase family of proteins, which are two-component flavin-dependent oxygenases (Li et al., 2008). LadA is believed to oxidize alkanes by a mechanism similar to that of other flavoprotein monooxygenases, so that its ability to recognize and hydroxylate long-chain length alkanes probably lies in the way it captures the alkane. Several bacterial strains can degrade >C20 alkanes using enzyme systems that have still not been characterized. It is likely that new enzyme classes will be found in the near future responsible for the oxidation of these high molecular weight alkanes.

4.5

Several Alkane Hydroxylases Frequently Coexist in a Single Bacterial Strain

Some bacterial strains contain only one alkane hydroxylase, as is the case for the wellcharacterized alkane degrader P. putida GPo1. However, it is rather common to find strains that contain more than one alkane oxidation system. In many cases, but not always, these alkane oxidation enzymes have different substrate ranges or different induction patterns. A. borkumensis SK2, which is specialized in assimilating alkanes, has two AlkB-related alkane hydroxylases and two genes encoding cytochrome P450s believed to be involved in alkane degradation, one of which is duplicated (Hara et al., 2004; Sabirova et al., 2006; Schneiker et al., 2006; van Beilen et al., 2004). As mentioned above, two genes that may encode alkane hydroxylases similar to AlmA were also detected in this bacterium (Throne-Holst et al., 2007). The presence of multiple alkane oxidation determinants in a single strain is not restricted to hydrocarbonoclastic bacteria, and can be observed as well in bacterial species that have a versatile metabolism. P. aeruginosa strains PAO1 and RR1 contain two AlkB-related alkane hydroxylases that are differentially regulated (Marı´n et al., 2001; Stover et al., 2000). Acinetobacter sp. DSM17874, and probably other Acinetobacter strains, have at least three alkane oxidation enzymes, two of them involved in the degradation of C10–C20 alkanes and a third one that oxidizes C32–C36 alkanes (Throne-Holst et al., 2007). Acinetobacter sp. M1, besides the two AlkB-related hydroxylases, contains a dioxygenase that oxidizes long-chainlength alkanes (Maeng et al., 1996; Tani et al., 2001) and a gene coding for a protein similar to AlmA (Throne-Holst et al., 2007). Mycobacterium sp. TY-6 and Nocardioides sp. CF8 also contain two different alkane oxidation systems for alkanes of different size ranges (Hamamura et al., 2001; Kotani et al., 2006). Rhodococcus strains Q15 and NRRL B-16531 contain at least four AlkB-related alkane hydroxylases (Whyte et al., 2002) and, in the latter strain, two additional cytochrome P450s of the CYP153 family have been detected (van Beilen et al., 2006). It is clear, therefore, that the coexistence of several alkane degradation systems is not uncommon. The presence of different and frequently highly divergent alkane degradation genes in a single bacterial strain suggests that horizontal transfer has greatly facilitated the spread of these genes. A phylogenetic analysis of 58 AlkB-related proteins identified in different Gram-positive and Gram-negative bacteria showed that AlkB homologs from fluorescent Pseudomonads were almost as divergent as the entire set of genes analyzed (van Beilen et al., 2003). Similarly, the four AlkB-related proteins present in Rhodococcus strains Q15 and NRRL B-16531 are

Enzymes for Aerobic Degradation of Alkanes

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as divergent as all hydroxylases analyzed from Gram-positive strains (Whyte et al., 2002). Some alkane degradation genes have been found on transposons (van Beilen et al., 2001) or on plasmids (Sekine et al., 2006; van Beilen et al., 1994), which clearly facilitates their horizontal transfer. It is worth noting that the two AlkB genes present in A. borkumensis SK2 are located in two separate genome islands that were probably acquired from an ancestor of the Yersinia lineage, and lately transferred from Alkanivorax to Pseudomonas (Reva et al., 2008).

5

Metabolism of the Alcohols and Aldehydes Derived from the Oxidation of Alkanes

The primary fatty alcohols generated by terminal oxidation of alkanes are further oxidized to aldehydes by an alcohol dehydrogenase (ADH). There are several kinds of ADHs. Some use NAD(P)+ as electron acceptor, while others do not depend on NAD(P)+ and use electron acceptors such as cytochromes or ubiquinone. Most NAD(P)+-independent ADHs contain pyrroloquinoline quinone (PQQ) as prosthetic group, and are commonly named quinoprotein ADHs. Many bacteria contain several different ADHs that can be used for the assimilation of distinct alcohols. For example, P. butanovora can express at least four different ADHs with different specificities towards primary and secondary alcohols (Vangnai and Arp, 2001; Vangnai et al., 2002). Assimilation of the alcohols derived from butane relies on two NAD+independent primary ADHs, named BDH and BOH (Vangnai and Arp, 2001; Vangnai et al., 2002). BDH contains PQQ and heme c as prostetic groups, while BOH contains only PQQ. Both enzymes recognize a broad range of substrates; BDH oxidizes C2–C8 primary alcohols, C5–C9 secondary alcohols and several aldehydes (Vangnai and Arp, 2001), while BOH is active on C2–C8 primary alcohols and C3–C8 secondary alcohols (Vangnai et al., 2002). Growth of cells in butane leads to induction of the genes coding for these two enzymes. Insertional inactivation of the gene coding for BDH, or of that coding for BOH, impairs but does not eliminate assimilation of butane, although the simultaneous inactivation of both genes renders cells unable to grow on butane (Vangnai et al., 2002). When P. butanovora is grown on 2-butanol and lactate, two additional NAD+-dependent secondary ADHs have been detected, although their role has not been analyzed in detail (Vangnai and Arp, 2001). The aldehydes generated by BOH and BDH are further oxidized to fatty acids. Genes coding for enzymes showing similarity to aldehyde dehydrogenases have been observed next to those coding for BOH and BDH, although their precise role has not been reported (Vangnai et al., 2002). It is worth noting that BOH and BDH are active towards aldehydes (Vangnai and Arp, 2001; Vangnai et al., 2002). Acinetobacter calcoaceticus HO1-N contains at least two ADHs. One of them requires NAD+ and shows preference for decanol. The other one requires NADP+ and has higher activity towards tetradecanol. An aldehyde dehydrogenase active towards long-chain aldehydes has also been described in this strain (Fox et al., 1992; Singer and Finnerty, 1985a, b), as well as in Acinetobacter sp. M1 (Ishige et al., 2000). Genes coding for alcohol and aldehyde dehydrogenases are also present in the in the P. putida GPo1 OCT plasmid. The alcohol dehydrogenase, named AlkJ, is necessary for growth on n-alkanes only if the chromosomal AlcA alcohol dehydrogenase is inactivated by mutation (van Beilen et al., 1992a), indicating again a redundancy in these enzymes. Similarly, the plasmid encoded AlkH aldehyde dehydrogenase is not essential for growth on alkanes, which

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agrees with the presence of several aldehyde dehydrogenases in the P. putida GPo1 chromosome (van Beilen et al., 1994). In the case of the secondary alcohols generated by subterminal oxidation of alkanes, alcohol dehydrogenases transform them into ketones (> Fig. 1). Gordonia sp. strain TY-5, a bacterium that can grow at the expense of propane and C13–C22 alkanes, metabolizes propane via 2-propanol and contains three NAD+-dependent secondary ADHs (Kotani et al., 2003). Although 2-propanol can be oxidized by any of the three secondary ADHs, which are all expressed in propane-grown cells, ADH1 seemed to play the major role under the conditions analyzed. NAD+-dependent secondary ADHs have been identified in other bacteria such as R. rhodochrous PNKb1 (Ashraf and Murrell, 1990), M. vaccae JOB5 (Coleman and Perry, 1985) and P. fluorescens NRRL B-1244 (Hou et al., 1983). The fatty acids generated by oxidation of the aldehydes are further metabolized by b-oxidation, generating Acyl-CoA that enters the tricarboxylic acids cycle. However, when the carbon source is in excess relative to nitrogen, many bacteria derive part of the carbon to generate storage materials such as triacylglycerols, wax esters, poly(hydroxybutyrate) or poly (3-hydroxyalkanoates), which accumulate as lipid bodies or as granules (Alvarez and Steinbuchel, 2002; Prieto, 2007; Waltermann et al., 2005). These compounds can later serve as endogenous carbon and energy sources during starvation periods. Formation of storage lipids is frequent among hydrocarbon utilizing-marine bacteria. Alcanivorax strains, for example, can accumulate triacylglycerols and wax esters when growing at the expense of pyruvate or n-alkanes (Kalscheuer et al., 2007). On the other hand, P. putida GPo1, a soil bacterium, can form intracellular inclusions of poly-b-hydroxyoctanoate when grown on n-octane (de Smet et al., 1983), while Acinetobacter sp. M-1 forms wax esters when growing at the expense of hexadecane (Ishige et al., 2000, 2002).

6

Degradation of Branched-Chain Alkanes

Branched-chain alkanes are more difficult to degrade than linear n-alkanes. It was observed long ago that n-alkanes are preferentially assimilated over branched alkanes (Pirnik et al., 1974). However, several bacterial strains can degrade simple branched-chain alkanes such as isooctane (Solano-Serena et al., 2004), or much more complex compounds like pristane (reviewed in Britton, 1984; Watkinson and Morgan, 1990). Alcanivorax sp. can also degrade branched alkanes such as pristane and phytane, a property that seems to provide a competitive advantage in oil-contaminated sea water (Hara et al., 2003). The metabolic pathways responsible for the assimilation of branched alkanes are less well characterized than those for n-alkanes, and may involve an o- or b-oxidation of the hydrocarbon molecule (Watkinson and Morgan, 1990).

7

Applications of Alkane Oxidation Enzymes in Biotransformations of Industrial Interest

In addition to their role in alkane degradation, alkane hydroxylases can be useful in biotransformation processes since they frequently oxidize not only their natural substrates, but other compounds as well, albeit with reduced efficiency. P. putida GPo1 AlkB can, for example, generate epoxides from alkenes and other chemicals with a terminal double bond, it oxidizes

Enzymes for Aerobic Degradation of Alkanes

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alcohols to aldehydes and catalyzes demethylation and sulfoxidation reactions (van Beilen et al., 1996; Witholt et al., 1990). It can also oxidize methyl tert-butyl ether (Smith and Hyman, 2004). Oxidation is regio- and stereo-specific which, in the case of some substrates, opens doors for applications in fine chemistry. For example, when acting on a compound with a terminal double bond it produces an (R)-epoxide with high enantiomeric excess. Optically active epoxides can be used to generate a number of chemicals that are useful precursors from which to derive several value-added products. The set-up of a cost-effective high-scale process based on this enzyme is complicated, however, due to practical issues such as substrate uptake, toxicity of the substrate and/or the product generated, uncoupling, oxygen mass transfer, low turnover with some compounds, or problems related to product recovery.

8

Research Needs

In spite of the extensive research on alkane degradation by bacteria performed during several decades, there are still aspects that remain poorly understood. One is how alkanes are incorporated or transported into the cell, which may differ for different alkanes and for different microorganisms. The enzymes for the degradation of low- and medium-chain length alkanes are rather well characterized, except for the paucity of structural data. However, some findings indicate that C20–C50 alkanes are probably oxidized by enzymes that have still not been identified. It is also rather intriguing why bacterial strains frequently contain several different or related alkane hydroxylases that have very similar substrate specificities. It may be that these hydroxylases differ in aspects that are still unknown but that are important for cell biology. Finally, the use of alkane hydroxylases for biotransformations of industrial interest, which has a great potential, still has to solve several technical issues that limit the efficiency of the process.

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Enzymes for Aerobic Degradation of Alkanes cellular inclusions and role of acyl coenzyme A reductase. Appl Environ Microbiol 68: 1192–1195. Johnson EL, Hyman MR (2006) Propane and n-butane oxidation by Pseudomonas putida GPo1. Appl Environ Microbiol 72: 950–952. Kalscheuer R, Stoveken T, Malkus U, Reichelt R, Golyshin PN, Sabirova JS, Ferrer M, Timmis KN, Steinbuchel A (2007) Analysis of storage lipid accumulation in Alcanivorax borkumensis: evidence for alternative triacylglycerol biosynthesis routes in bacteria. J Bacteriol 189: 918–928. Kasai Y, Kishira H, Sasaki T, Syutsubo K, Watanabe K, Harayama S (2002) Predominant growth of Alcanivorax strains in oil-contaminated and nutrientsupplemented sea water. Environ Microbiol 4: 141–147. Kohler T, Curty LK, Barja F, van Delden C, Pechere JC (2000) Swarming of Pseudomonas aeruginosa is dependent on cell-to-cell signaling and requires flagella and pili. J Bacteriol 182: 5990–5996. Kotani T, Kawashima Y, Yurimoto H, Kato N, Sakai Y (2006) Gene structure and regulation of alkane monooxygenases in propane-utilizing Mycobacterium sp. TY-6 and Pseudonocardia sp. TY-7. J Biosci Bioeng 102: 184–192. Kotani T, Yamamoto T, Yurimoto H, Sakai Y, Kato N (2003) Propane monooxygenase and NAD+dependent secondary alcohol dehydrogenase in propane metabolism by Gordonia sp. strain TY-5. J Bacteriol 185: 7120–7128. Kotani T, Yurimoto H, Kato N, Sakai Y (2007) Novel acetone metabolism in a propane-utilizing bacterium, Gordonia sp. strain TY-5. J Bacteriol 189: 886–893. Kurth EG, Doughty DM, Bottomley PJ, Arp DJ, Sayavedra-Soto LA (2008) Involvement of BmoR and BmoG in n-alkane metabolism in ‘‘Pseudomonas butanovora.’’ Microbiology 154: 139–147. Li L, Liu X, Yang W, Xu F, Wang W, Feng L, Bartlam M, Wang L, Rao Z (2008) Crystal structure of longchain alkane monooxygenase (LadA) in complex with coenzyme FMN: unveiling the long-chain alkane hydroxylase. J Mol Biol 376: 453–465. Maeng JH, Sakai Y, Ishige T, Tani Y, Kato N (1996) Diversity of dioxygenases that catalyze the first step of oxidation of long-chain n-alkanes in Acinetobacter sp. strain M-1. FEMS Microbiol Lett 141: 177–182. Maier T, Forster HH, Asperger O, Hahn U (2001) Molecular characterization of the 56-kDa CYP153 from Acinetobacter sp. EB104. Biochem Biophys Res Commun 286: 652–658. Margesin R, Labbe D, Schinner F, Greer CW, Whyte LG (2003) Characterization of hydrocarbon-degrading microbial populations in contaminated and pristine Alpine soils. Appl Environ Microbiol 69: 3085–3092.

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Marı´n MM, Smits TH, van Beilen JB, Rojo F (2001) The alkane hydroxylase gene of Burkholderia cepacia RR10 is under catabolite repression control. J Bacteriol 183: 4202–4209. Marı´n MM, Yuste L, Rojo F (2003) Differential expression of the components of the two alkane hydroxylases from Pseudomonas aeruginosa. J Bacteriol 185: 3232–3237. McKew BA, Coulon F, Osborn AM, Timmis KN, McGenity TJ (2007a) Determining the identity and roles of oil-metabolizing marine bacteria from the Thames estuary, UK. Environ Microbiol 9: 165–176. McKew BA, Coulon F, Yakimov MM, Denaro R, Genovese M, Smith CJ, Osborn AM, Timmis KN, McGenity TJ (2007b) Efficacy of intervention strategies for bioremediation of crude oil in marine systems and effects on indigenous hydrocarbonoclastic bacteria. Environ Microbiol 9: 1562–1571. Neu TR (1996) Significance of bacterial surface-active compounds in interaction of bacteria with interfaces. Microbiol Rev 60: 151–166. Noordman WH, Janssen DB (2002) Rhamnolipid stimulates uptake of hydrophobic compounds by Pseudomonas aeruginosa. Appl Environ Microbiol 68: 4502–4508. Ohkuma M, Zimmer T, Iida T, Schunck WH, Ohta A, Takagi M (1998) Isozyme function of n-alkane-inducible cytochromes P450 in Candida maltosa revealed by sequential gene disruption. J Biol Chem 273: 3948–3953. Pirnik MP, Atlas RM, Bartha R (1974) Hydrocarbon metabolism by Brevibacterium erythrogenes: normal and branched alkanes. J Bacteriol 119: 868–878. Prieto MA (2007) From oil to bioplastics, a dream come true? J Bacteriol 189: 289–290. Rehm HJ, Reiff I (1981) Mechanisms and occurrence of microbial oxidation of long-chain alkanes. Adv Biochem Eng 19: 175–215. Reva ON, Hallin PF, Willenbrock H, Sicheritz-Ponten T, Tu¨mmler B, Ussery DW (2008) Global features of the Alcanivorax borkumensis SK2 genome. Environ Microbiol 10: 614–625. Rojo F (2005) Specificity at the end of the tunnel: understanding substrate length discrimination by the AlkB alkane hydroxylase. J Bacteriol 187: 19–22. Ron EZ, Rosenberg E (2001) Natural roles of biosurfactants. Environ Microbiol 3: 229–236. Ron EZ, Rosenberg E (2002) Biosurfactants and oil bioremediation. Curr Opin Biotechnol 13: 249–252. Rosenberg E (1993) Exploiting microbial growth on hydrocarbons – new markets. Trends Biotechnol 11: 419–424. Sabirova JS, Ferrer M, Regenhardt D, Timmis KN, Golyshin PN (2006) Proteomic insights into metabolic adaptations in Alcanivorax borkumensis induced by alkane utilization. J Bacteriol 188: 3763–3773.

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Schmitz C, Goebel I, Wagner S, Vomberg A, Klinner U (2000) Competition between n-alkane-assimilating yeasts and bacteria during colonization of sandy soil microcosms. Appl Microbiol Biotechnol 54: 126–132. Schneiker S, Martins dos Santos VA, Bartels D, Bekel T, Brecht M, Buhrmester J, Chernikova TN, Denaro R, Ferrer M, Gertler C, Goesmann A, Golyshina OV, Kaminski F, Khachane AN, Lang S, Linke B, McHardy AC, Meyer F, Nechitaylo T, Puhler A, Regenhardt D, Rupp O, Sabirova JS, Selbitschka W, Yakimov MM, Timmis KN, Vorholter FJ, Weidner S, Kaiser O, Golyshin PN (2006) Genome sequence of the ubiquitous hydrocarbon-degrading marine bacterium Alcanivorax borkumensis. Nat Biotechnol 24: 997–1004. Sekine M, Tanikawa S, Omata S, Saito M, Fujisawa T, Tsukatani N, Tajima T, Sekigawa T, Kosugi H, Matsuo Y, Nishiko R, Imamura K, Ito M, Narita H, Tago S, Fujita N, Harayama S (2006) Sequence analysis of three plasmids harboured in Rhodococcus erythropolis strain PR4. Environ Microbiol 8: 334–346. Shanklin J, Achim C, Schmidt H, Fox BG, Munck E (1997) Mossbauer studies of alkane omegahydroxylase: evidence for a diiron cluster in an integral-membrane enzyme. Proc Natl Acad Sci USA 94: 2981–2986. Shanklin J, Whittle E, Fox BG (1994) Eight histidine residues are catalytically essential in a membraneassociated iron enzyme, stearoyl-CoA desaturase, and are conserved in alkane hydroxylase and xylene monooxygenase. Biochemistry 33: 12787–12794. Singer ME, Finnerty WR (1985a) Alcohol dehydrogenases in Acinetobacter sp. strain HO1-N: role in hexadecane and hexadecanol metabolism. J Bacteriol 164: 1017–1024. Singer ME, Finnerty WR (1985b) Fatty aldehyde dehydrogenases in Acinetobacter sp. strain HO1-N: role in hexadecanol metabolism. J Bacteriol 164: 1011–1016. Sluis MK, Sayavedra-Soto LA, Arp DJ (2002) Molecular analysis of the soluble butane monooxygenase from ‘‘Pseudomonas butanovora.’’ Microbiology 148: 3617–3629. Smith CA, Hyman MR (2004) Oxidation of methyl tert-butyl ether by alkane hydroxylase in dicyclopropylketone-induced and n-octane-grown Pseudomonas putida GPo1. Appl Environ Microbiol 70: 4544–4550. Smits TH, Balada SB, Witholt B, van Beilen JB (2002) Functional analysis of alkane hydroxylases from Gram-negative and Gram-positive bacteria. J Bacteriol 184: 1733–1742. Smits TH, Witholt B, van Beilen JB (2003) Functional characterization of genes involved in alkane oxidation by Pseudomonas aeruginosa. Antonie Van Leeuwenhoek 84: 193–200.

Smits THM, Ro¨thlisberger M, Witholt B, van Beilen JB (1999) Molecular screening for alkane hydroxylase genes in Gram-negative and Gram-positive strains. Environ Microbiol 1: 307–317. Solano-Serena F, Marchal R, Heiss S, Vandecasteele JP (2004) Degradation of isooctane by Mycobacterium austroafricanum IFP 2173: growth and catabolic pathway. J Appl Microbiol 97: 629–639. Stover CK, Pham XQ, Erwin AL, Mizoguchi SD, Warrener P, Hickey MJ, Brinkman FS, Hufnagle WO, Kowalik DJ, Lagrou M, Garber RL, Goltry L, Tolentino E, Westbrock-Wadman S, Yuan Y, Brody LL, Coulter SN, Folger KR, Kas A, Larbig K, Lim R, Smith K, Spencer D, Wong GK, Wu Z, Paulsen IT (2000) Complete genome sequence of Pseudomonas aeruginosa PA01, an opportunistic pathogen. Nature 406: 959–964. Tani A, Ishige T, Sakai Y, Kato N (2001) Gene structures and regulation of the alkane hydroxylase complex in Acinetobacter sp. strain M-1. J Bacteriol 183: 1819–1823. Throne-Holst M, Markussen S, Winnberg A, Ellingsen TE, Kotlar HK, Zotchev SB (2006) Utilization of n-alkanes by a newly isolated strain of Acinetobacter venetianus: the role of two AlkB-type alkane hydroxylases. Appl Microbiol Biotechnol 72: 353–360. Throne-Holst M, Wentzel A, Ellingsen TE, Kotlar HK, Zotchev SB (2007) Identification of novel genes involved in long-chain n-alkane degradation by Acinetobacter sp. strain DSM 17874. Appl Environ Microbiol 73: 3327–3332. van Beilen JB, Eggink G, Enequist H, Bos R, Witholt B (1992a) DNA sequence determination and functional characterization of the OCT-plasmidencoded alkJKL genes of Pseudomonas oleovorans. Mol Microbiol 6: 3121–3136. van Beilen JB, Funhoff EG (2007) Alkane hydroxylases involved in microbial alkane degradation. Appl Microbiol Biotechnol 74: 13–21. van Beilen JB, Funhoff EG, van Loon A, Just A, Kaysser L, Bouza M, Holtackers R, Rothlisberger M, Li Z, Witholt B (2006) Cytochrome P450 alkane hydroxylases of the CYP153 family are common in alkanedegrading eubacteria lacking integral membrane alkane hydroxylases. Appl Environ Microbiol 72: 59–65. van Beilen JB, Holtackers R, Luscher D, Bauer U, Witholt B, Duetz WA (2005a) Biocatalytic production of perillyl alcohol from limonene by using a novel Mycobacterium sp. cytochrome P450 alkane hydroxylase expressed in Pseudomonas putida. Appl Environ Microbiol 71: 1737–1744. van Beilen JB, Li Z, Duetz WA, Smits THM, Witholt B (2003) Diversity of alkane hydroxylase systems in the environment. Oil Gas Sci Technol 58: 427–440.

Enzymes for Aerobic Degradation of Alkanes van Beilen JB, Marin MM, Smits TH, Rothlisberger M, Franchini AG, Witholt B, Rojo F (2004) Characterization of two alkane hydroxylase genes from the marine hydrocarbonoclastic bacterium Alcanivorax borkumensis. Environ Microbiol 6: 264–273. van Beilen JB, Neuenschwander M, Smits TH, Roth C, Balada SB, Witholt B (2002a) Rubredoxins involved in alkane oxidation. J Bacteriol 184: 1722–1732. van Beilen JB, Panke S, Lucchini S, Franchini AG, Ro¨thlisberger M, Witholt B (2001) Analysis of Pseudomonas putida alkane degradation gene clusters and flanking insertion sequences: evolution and regulation of the alk-genes. Microbiology 147: 1621–1630. van Beilen JB, Penninga D, Witholt B (1992b) Topology of the membrane-bound alkane hydroxylase of Pseudomonas oleovorans. J Biol Chem 267: 9194–9201. van Beilen JB, Smits TH, Roos FF, Brunner T, Balada SB, Rothlisberger M, Witholt B (2005b) Identification of an amino acid position that determines the substrate range of integral membrane alkane hydroxylases. J Bacteriol 187: 85–91. van Beilen JB, Smits TH, Whyte LG, Schorcht S, Rothlisberger M, Plaggemeier T, Engesser KH, Witholt B (2002b) Alkane hydroxylase homologues in Gram-positive strains. Environ Microbiol 4: 676–682. van Beilen JB, Wubbolts MG, Chen Q, Nieboer M, Witholt B (1996) Effects of two-liquid-phase systems and expression of alk genes on the physiology of alkane-oxidizing strains. In Molecular Biology of Pseudomonads. T Nakazawa, K Furukawa, D Haas, and S Silver (eds.). Washington, DC: ASM Press, pp. 35–47. van Beilen JB, Wubbolts MG, Witholt B (1994) Genetics of alkane oxidation by Pseudomonas oleovorans. Biodegradation 5: 161–174. Vangnai AS, Arp DJ (2001) An inducible 1-butanol dehydrogenase, a quinohaemoprotein, is involved in the oxidation of butane by ‘‘Pseudomonas butanovora.’’ Microbiology 147: 745–756. Vangnai AS, Arp DJ, Sayavedra-Soto LA (2002) Two distinct alcohol dehydrogenases participate in butane metabolism by Pseudomonas butanovora. J Bacteriol 184: 1916–1924. van Hamme JD, Singh A, Ward OP (2003) Recent advances in petroleum microbiology. Microbiol Mol Biol Rev 67: 503–549. Waltermann M, Hinz A, Robenek H, Troyer D, Reichelt R, Malkus U, Galla HJ, Kalscheuer R, Stoveken T, von Landenberg P, Steinbuchel A (2005) Mechanism of lipid-body formation in prokaryotes: how bacteria fatten up. Mol Microbiol 55: 750–763.

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Watkinson RJ, Morgan P (1990) Physiology of aliphatic hydrocarbon-degrading microorganisms. Biodegradation 1: 79–92. Wentzel A, Ellingsen TE, Kotlar HK, Zotchev SB, Throne-Holst M (2007) Bacterial metabolism of long-chain n-alkanes. Appl Microbiol Biotechnol 76: 1209–1221. Whyte LG, Hawari J, Zhou E, Bourbonniere L, Inniss WE, Greer CW (1998) Biodegradation of variable-chain-length alkanes at low temperatures by a psychrotrophic Rhodococcus sp. Appl Environ Microbiol 64: 2578–2584. Whyte LG, Smits TH, Labbe D, Witholt B, Greer CW, Van Beilen JB (2002) Gene cloning and characterization of multiple alkane hydroxylase systems in Rhodococcus strains Q15 and NRRL B-16531. Appl Environ Microbiol 68: 5933–5942. Witholt B, de Smet MJ, Kingma J, van Beilen JB, Kok M, Lageveen RG, Eggink G (1990) Bioconversions of aliphatic compounds by Pseudomonas oleovorans in multiphase bioreactors: background and economic potential. Trends Biotechnol 8: 46–52. Yakimov MM, Giuliano L, Denaro R, Crisafi E, Chernikova TN, Abraham WR, Luensdorf H, Timmis KN, Golyshin PN (2004) Thalassolituus oleivorans gen. nov., sp. nov., a novel marine bacterium that obligately utilizes hydrocarbons. Int J Syst Evol Microbiol 54: 141–148. Yakimov MM, Giuliano L, Gentile G, Crisafi E, Chernikova TN, Abraham WR, Lunsdorf H, Timmis KN, Golyshin PN (2003) Oleispira antarctica gen. nov., sp. nov., a novel hydrocarbonoclastic marine bacterium isolated from Antarctic coastal sea water. Int J Syst Evol Microbiol 53: 779–785. Yakimov MM, Golyshin PN, Lang S, Moore ER, Abraham WR, Lunsdorf H, Timmis KN (1998) Alcanivorax borkumensis gen. nov., sp. nov., a new, hydrocarbon-degrading and surfactant-producing marine bacterium. Int J Syst Bacteriol 48(Pt 2): 339–348. Yakimov MM, Timmis KN, Golyshin PN (2007) Obligate oil-degrading marine bacteria. Curr Opin Biotechnol 18: 257–266. Zimmer T, Ohkuma M, Ohta A, Takagi M, Schunck WH (1996) The CYP52 multigene family of Candida maltosa encodes functionally diverse n-alkane-inducible cytochromes P450. Biochem Biophys Res Commun 224: 784–789. Zulianello L, Canard C, Kohler T, Caille D, Lacroix JS, Meda P (2006) Rhamnolipids are virulence factors that promote early infiltration of primary human airway epithelia by Pseudomonas aeruginosa. Infect Immun 74: 3134–3147.

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4 Aerobic Degradation of Aromatic Hydrocarbons D. Pe´rez-Pantoja1 . B. Gonza´lez1 . D. H. Pieper2,* 1 Departamento de Gene´tica Molecular y Microbiologı´a, Facultad de Ciencias Biolo´gicas, NM-EMBA, CASEB, P. Universidad Cato´lica de Chile, Santiago, Chile 2 Biodegradation Research Group, Department of Microbial Pathogenesis, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 800

2 2.1

2.4

Catabolism of Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 800 Peripheral Reactions Preparing Aromatic Hydrocarbons for Ring-Cleavage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 801 Rieske Non-Heme Iron Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 801 Soluble Diiron Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 804 Single and Two Component Flavoprotein Monooxygenases . . . . . . . . . . . . . . . . . . . . . . 805 Side Chain Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 808 Oxidations of Methyl Groups in Methyl-Substituted Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 808 O-Demethylations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 810 Aromatic Acid Decarboxylations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 810 CoA-Dependent Peripheral Pathways for Phenylpropenoid Compounds . . . . . . . . . 812 Peripheral Reactions in the Degradation of Aromatic Amino Acids . . . . . . . . . . . . . . 814 Central Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 814 Intradiol Ring-Cleavage Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 814 Catechol Meta-Cleavage Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 817 Protocatechuate Meta-Cleavage Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 818 Further Meta-Cleavage Routes Involving Type I or Type II Extradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 821 Pathways Involving Extradiol Ring-Cleavage by Enzymes of the Cupin Superfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 823 CoA Dependent Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 826

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 829

2.1.1 2.1.2 2.1.3 2.2 2.2.1 2.2.2 2.2.3 2.2.4 2.2.5 2.3 2.3.1 2.3.2 2.3.3 2.3.4 2.3.5

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_60, # Springer-Verlag Berlin Heidelberg, 2010

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Aerobic Degradation of Aromatic Hydrocarbons

Abstract: Aromatic hydrocarbons are widely distributed in nature. They are found as lignin components, aromatic amino acids and xenobiotic compounds, among others. Microorganisms, mostly bacteria, degrade an impressive variety of such chemical structures. The major principle of aromatic hydrocarbon biodegradation is that a broad range of peripheral reactions are transformed to a restricted range of central intermediates, which are subject to ringcleavage and funneling into the Krebs cycle. Key enzymes in aerobic aromatic degradation are oxygenases, preparing aromatics for ring-cleavage by the introduction of hydroxyl functions and catalyzing cleavage of the aromatic ring. The diverse monooxygenases and dioxygenases involved in hydroxylations, a significant proportion of them possessing relaxed substrate specificity, are discussed as well as the broad diversity of side chain processing transformations involved in the formation of ring-cleavage central intermediates. Ring cleavage dioxygenases, covering intradiol ring cleavage of ortho dihydroxylated intermediates, and a large number of diverse but mechanistically related extradiol dioxygenases participating in ring cleavage of ortho and para dihydroxylated intermediates are also discussed. Novel CoA dependent aerobic routes to allow ring-cleavage of aromatic hydrocarbons without involvement of dihydroxylated aromatic intermediates have been described in the last years and are also reviewed. The degradation of heteroarenes will not be described in this chapter.

1

Introduction

Aromatic hydrocarbons are very important building blocks of biomass and are widely distributed in nature, being produced by a variety of biological and biogeochemical processes, and range in size from low-molecular mass compounds to polymers. The most abundant fraction of aromatic hydrocarbons is formed by the lignin of higher plants, which in fact is the second most abundant polymer in nature after cellulose, comprising about 25% of the land-based biomass on Earth (Kirk and Farrell, 1987). During the decomposition process, lignin degradation products – together with other plant-derived aromatic hydrocarbons – contribute to the formation of recalcitrant organic matter in soils. Other ubiquitous sources of aromatic hydrocarbons are the aromatic amino acids. Finally, the extensive use of natural and xenobiotic aromatic hydrocarbons in industrial processes, coupled with inadequate waste management strategies, has led to the positioning of these compounds among the most stable and persistent organic pollutants. The degradation of aromatic polymers is an important component of global biogeochemical cycles and is accomplished almost exclusively by microorganisms which have evolved diverse strategies to degrade aromatic hydrocarbons, and thereby derive carbon and energetic benefit from them. The large spectrum of aromatic substrates degradable by microbial communities is assured by a huge catabolic diversity present in different microbial species and the relaxed substrate specificity of some of the catabolic pathways (Pe´rez-Pantoja et al., 2008, > Chapter 39, Vol. 2, Part 6). The major principle of aromatic hydrocarbon biodegradation is that catabolic pathways involve two key steps: the activation of the thermodynamically stable benzene ring from structurally diverse aromatics, and its subsequent cleavage. In aerobic catabolism, oxygenases accomplish the main role in both steps.

2

Catabolism of Aromatic Hydrocarbons

Aerobic microorganisms usually initiate degradation by activation of the aromatic nucleus through oxygenation reactions. A few central intermediates such as catechols, protocatechuates, gentisates and (hydroxy)benzoquinols, are produced by the introduction of hydroxyl

Aerobic Degradation of Aromatic Hydrocarbons

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groups, usually in ortho- or para-position to one another. These intermediates are subject to oxygenolytic ring cleavage followed by channeling of the ring-cleavage products into the central metabolism. Alternatively aromatic hydrocarbons, even under aerobic conditions, can be metabolized through the corresponding CoA thioesters and subject of non-oxygenolytic ring cleavage.

2.1

Peripheral Reactions Preparing Aromatic Hydrocarbons for Ring-Cleavage

2.1.1

Rieske Non-Heme Iron Oxygenases

The so called Rieske non-heme iron oxygenases are one of the key families of enzymes important for aerobic activation and thus degradation of aromatic hydrocarbons such as benzoate, benzene, toluene, phthalate, naphthalene or biphenyl (Gibson and Parales, 2000). These multicomponent enzyme complexes, composed of a terminal oxygenase component and different electron transport proteins, usually catalyze the incorporation of two oxygen atoms into the aromatic ring to form arene-cis-dihydrodiols (although some members of this superfamily also catalyze monooxygenations), a reaction which is followed by a dehydrogenation usually catalyzed by cis-dihydrodiol dehydrogenases to give (substituted) catechols. Members of the Rieske non-heme iron oxygenases are since decades known to be involved in the degradation of benzoate (Gibson et al., 1968), converting it to 1-carboxy-1,2-cisdihydroxycyclohexa-3,5-diene (benzoate-cis-dihydrodiol, see > Fig. 1) (Reiner and Hegeman, 1971). A benzoate dihydrodiol dehydrogenase catalyzes the dehydrogenation to a b-ketoacid, which spontaneously decarboxylates to catechol (Reiner, 1972). The benzoate dioxygenases belong to the so-called benzoate subgroup of Rieske non-heme iron oxygenases and are composed of a reductase and an oxygenase component with an (ab)3 quaternary structure (Wolfe et al., 2002), with each a-subunit containing a mononuclear non-heme iron active site and a Rieske-type (2Fe-2S) cluster. Some of these benzoate dioxygenases have been studied in detail and it is well established that despite a significant degree in sequence identity, they differ significantly in substrate specificity (Reineke and Knackmuss, 1978), with toluate dioxygenase of P. putida mt-2 being capable to transform meta- and para-substituted benzoates whereas benzoate dioxygenase only transforms benzoate and meta-substituted benzoates. Ortho-substituted benzoates (2-chloro- and 2-methyl-) are poor substrates for both enzymes (Yamaguchi and Fujisawa, 1980). Similar two-component enzyme systems (see > Fig. 1) are responsible for 1,2-dioxygenation of anthranilate (Bundy et al., 1998), an intermediary metabolite of tryptophan degradation and a precursor for the Pseudomonas quinolone signal (Farrow and Pesci, 2007). Anthranilate dioxygenases catalyze the formation of catechol without requirement of a dehydrogenase due to spontaneous decarboxylation and deamination of 2-amino-1-carboxy1,2-cis-dihydroxycyclohexa-3,5-diene (anthranilate-cis-dihydrodiol, see > Fig. 1). At least the enzyme system of Acinetobacter baylyi ADP1 also transforms benzoate (Eby et al., 2001), however, other ortho-substituted benzoates are only poorly converted, differentiating two-component anthranilate 1,2-dioxygenases from two-component 2-halobenzoate 1,2dioxygenases (See > Chapter 5, Vol. 2, Part 2). Rieske non-heme iron oxygenases are also involved in the degradation of p-cymene (p-isopropyltoluene), a natural product identified in volatile oils from various plants. p-Cymene metabolism is initiated by oxidation of the methyl substituent with p-cumate as

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. Figure 1 Dendrogram showing the relatedness among oxygenase a-subunits of Rieske non-heme iron oxygenases. Reactions catalyzed by enzymes indicated are given to the exterior of the figure, together with subsequent reactions resulting in the formation of central intermediates, which are subject to ring-cleavage reaction. Unstable intermediates are shown in brackets.

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Aerobic Degradation of Aromatic Hydrocarbons

4

intermediate (Eaton, 1997), which is attacked by a two-component p-cumate dioxygenase (Eaton, 1996). In contrast to enzymes described above, p-cumate dioxygenases do not attack on the carboxysubstituted carbon atom but on the meta- and ortho- carbon atom to form 2,3dihydroxy-2,3-dihydro-p-cumate, followed by dehydrogenation to 2,3-dihydroxy-p-cumate (> Fig. 1). Also aniline (aminobenzene) degradation seems to be mediated by related two-component dioxygenases and the a-subunits of the aniline dioxygenase system share significant identity with those of benzoate dioxygenases (see > Fig. 1). However, in contrast to all enzyme systems described above, aniline dioxygenase consists of five protein components, all necessary for a functional enzyme (Fukumori and Saint, 1997). Based on sequence similarities, it is proposed that three of them function as the large and small subunit of aniline dioxygenase and reductase, respectively. Additional proteins show similarity to glutamine synthetase and glutamine amidotransferase, respectively, and maybe are involved in transfer of the amino group or release of ammonia (Liang et al., 2005). Interestingly, enzyme systems catalyzing the transformation of anthranilate (and also of 2-halobenzoate), only distantly related to those of the benzoate subgroup of Rieske non-heme iron oxygenases, have also been described (See > Chapter 5, Vol. 2, Part 2 for 2-halobenzoate 1,2-dioxygenases). Anthranilate dioxygenase of Burkholderia cepacia DBO1 is a threecomponent Rieske non-heme iron dioxygenase composed of a reductase, a ferredoxin and a two-subunit oxygenase which, besides anthranilate, also transforms salicylate (but not 2-chlorobenzoate) to catechol (Chang et al., 2003). Besides three component 2-halobenzoate 1,2-dioxygenases, the enzymes most closely related to three-component anthranilate dioxygenase have been characterized as salicylate 1-hydroxylases (see > Fig. 1). The occurrence of three component salicylate 1-hydroxylases, contrasting the previously known single component flavoprotein monooxygenases (see 2.1.3.1), was first reported in Sphingobium sp. strain P2, which synthesized three isoenzymes (Pinyakong et al., 2003). However, despite the similarity and identical products formed from salicylate, anthranilate dioxygenase from B. cepacia DBO1 and multicomponent salicylate 1-hydroxylases are quite distinct and whereas anthranilate dioxygenase catalyzes a dioxygenation of anthranilate, salicylate 1-hydroxylase catalyzes a monooxygenation of anthranilate with 2-aminophenol as product (Jouanneau et al., 2007). Phylogenetic analyses (see > Fig. 1) show that the a-subunits of three-component anthranilate dioxygenase, salicylate 1-hydroxylase and 2-halobenzoate 1,2-dioxygenase form a distinct group of enzymes together with salicylate 5-hydroxylases and terephthalate dioxygenases. Salicylate 5-hydroxylase, transforming salicylate into gentisate, has initially been reported in Ralstonia sp. U2 (Fuenmayor et al., 1998). The encoding genes were identified to be in the same operon as those of the naphthalene dioxygenase and it was shown that naphthalene dioxygenase and salicylate 5-hydroxylase share the chain for the transport of electrons (Zhou et al., 2002), a ferredoxin reductase and ferredoxin, as in three component oxygenases. As mentioned above the a-subunits of terephthalate dioxygenases belong to the same subgroup of enzymes, all of them involved in the metabolism of carboxylated aromatics (see > Fig. 1). Terephthalate dioxygenase catalyzes a 1,2-dioxygenation with 1,2-dihydroxy3,5-cyclohexadiene-1,4-dicarboxylate as product, which is dehydrogenated, as described above for benzoate dihydrodiol, to a b-ketoacid which spontaneously decarboxylates to protocatechuate (Schla¨fli et al., 1994). Terephthahlate dioxygenase, at least from C. testosteroni T-2, seems to be of restricted substrate specificity and neither attacks benzoate nor isophthalate or phthalate (Schla¨fli et al., 1994). In contrast to salicylate 1- and 5-hydroxylase or

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anthranilate dioxygenase, terephthalate dioxygenase seems to be active as a two-component dioxygenase consisting of a- and b-subunits of the oxygenase and a reductase (Sasoh et al., 2006). The degradation of phthalate has initially been described in B. cepacia (Batie et al., 1987). In Proteobacteria, phthalate is subject to 4,5-dioxygenation giving rise to the dihydrodiol which is dehydrogenated to 4,5-dihydroxyphthalate (> Fig. 1). As the attack does not involve a carboxylated carbon atom, no spontaneous decarboxylation is involved. Decarboxylation is catalyzed by a 4,5-dihydroxyphthalate decarboxylase yielding protocatechuate (> Fig. 1) (Lee et al., 1994). The oxygenase of phthalate 4,5-dioxygenase differs from all above described Rieske non-heme iron oxygenases as being composed only of a-subunits, a feature shared with carbazole dioxygenases, 3-chlorobenzoate 4,5-dioxygenase and isophthalate dioxygenase from C. testosteroni YZW-D (Wang et al., 1995), the only enzyme acting on isophthalate described thus far. Interestingly, phthalate 4,5-dioxygenases have only been described in Proteobacteria, whereas Actinobacteria such as Arthrobacter keyseri 12B degrade phthalate via 3,4-dioxygenation and through 3,4-dihydroxyphthalate, which is decarboxylated to protocatechuate (> Fig. 1) as common intermediate of both the 4,5- and 3,4-dioxygenolytic pathways (Eaton, 2001). In contrast to phthalate 4,5-dioxygenase, phthalate 3,4-dioxygenase is a three-component dioxygenase composed of a two-subunit terminal oxygenase, a ferredoxin and a ferredoxin reductase, with the a-subunit being closely related to a-subunits of naphthalene or phenanthrene dioxygenases from Actinobacteria (> Fig. 1). However, not only the oxygenase systems of phthalate 4,5- and phthalate 3,4-dioxygenase are different. cis-3,4-dihydro-3,4dihydroxyphthalate dehydrogenase belongs to the aldo/keto reductase superfamily (Eaton, 2001), and differs from cis-4,5-dihydro-4,5-dihydroxyphthalate dehydrogenase, which belongs to the GFO/IDH/MOCA family (Chang and Zylstra, 1998). In addition, 3,4-dihydroxyphthalate decarboxylases are unrelated to 4,5-dihydroxyphthalate decarboxylases. Also the catabolism of 3-phenylpropionate (and cinnamate) can be initiated by dioxygenation through the action of a Rieske non-heme iron dioxygenase (> Fig. 1). 3-Phenylpropionate dioxygenase, similarly to p-cumate dioxygenase, inserts oxygen into positions 2- and 3- of the aromatic ring yielding cis-3-(3-carboxyethyl)-3,5-cyclohexadiene-1,2-diol, followed by dehydrogenation through a dehydrogenase of the short chain alcohol dehydrogenase family to 2,3-dihydroxyphenylpropionate (or 2,3-dihydroxycinnamate) (Diaz et al., 2001). 3-Phenylpropionate 2,3-dioxygenases are related to the benzene/toluene/isopropylbenzene/ biphenyl subgroup (enzymes typically described to be capable to transform the aforementioned compounds) of Rieske non-heme iron oxygenases (> Fig. 1), important for degradation of hydrophobic substrates (> Chapter 26, Vol. 2, Part 5).

2.1.2

Soluble Diiron Monooxygenases

Enzymes capable to monooxygenate benzene/toluene to phenol/methylphenol and phenols to catechols belong to an evolutionary related family of soluble diiron 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 diiron center contained in each a-subunit. Theses monooxygenases are

Aerobic Degradation of Aromatic Hydrocarbons

4

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 four-component alkene/aromatic monooxygenases (Leahy et al., 2003). The four component alkene/aromatic monooxygenases comprise enzymes that oxidize non-hydroxylated compounds and their gene clusters usually encode a ferredoxin component (Leahy et al., 2003). Monooxygenases that hydroxylate toluene at all three possible positions, producing 2-methyl-, 3-methyl- or 4-methylphenol have been described (Olsen et al., 1994; Shields et al., 1989; Whited and Gibson, 1991). However, later analysis has shown that the toluene monooxygenase of R. pickettii PK01, which had been reported previously to hydroxylate toluene at the meta position, producing primarily 3-methylphenol (Olsen et al., 1994), hydroxylates toluene predominantly at the para position producing 4-methylphenol (Fishman et al., 2004). Some of the enzymes of this subfamily, like the toluene monooxygenases of P. stutzeri OX1 (Bertoni et al., 1998) or R. pickettii PK01, or toluene 4-monooxygenase of P. mendocina KR1 have been shown to oxidize also phenol and methylphenols to the respective catechols and even further to 1,2,3-trihydroxybenzene (Tao et al., 2004b). The phenol hydroxylases comprise the multicomponent phenol hydroxylase of the methylphenol degrading Pseudomonas sp. strain CF600 (Shingler et al., 1992) and P. putida 35X (Ng et al., 1994) but also the toluene 2-monooxygenase of B. cepacia G4 (Newman and Wackett, 1995), among others. The respective gene clusters usually lack a ferredoxin gene (Leahy et al., 2003). Whereas all these monooxygenases share the capability to hydroxylate phenol and methylsubstituted derivatives, only a few enzymes of this group have been shown to hydroxylate the unactivated benzene nucleus. These enzymes, among them toluene 2-monooxygenase of strain G4, sequentially oxidize toluene to 2-methylphenol and further to 3-methylcatechol (Newman and Wackett, 1995). Phenol hydroxylase of P. stutzeri OX1 was also shown to be capable to transform benzene or toluene, however, the specificity constant kcat/Km was 2–3 orders of magnitude lower compared to phenol as substrate, evidencing phenol to be the highly preferred substrate (Cafaro et al., 2004). It should be noted that in the recent years, the crystal structure of toluene monooxygenase from P. stutzeri OX1 has been solved and various mutagenesis studies on this group of enzymes have been performed, inter alia to elucidate amino acid residues determining regioselectivity (Fishman et al., 2005; Tao et al., 2004a), but also to identify and change residues critical for phenol hydroxylation (Tao et al., 2004b; Vardar and Wood, 2004).

2.1.3

Single and Two Component Flavoprotein Monooxygenases

Flavoprotein monooxygenases are involved in a wide variety of biological processes including biosynthesis of antibiotics and siderophores or biodegradation of aromatic hydrocarbons. The reactions use NAD(P)H and O2 as co-substrates and insert one atom of oxygen into the substrate. These enzymes utilize a general cycle in which NAD(P)H reduces the flavin, and the reduced flavin reacts with O2 to form a C4a-(hydro)peroxyflavin intermediate, which is the oxygenating agent. Hydroxylation of the substrate yields the flavin-C4a-hydroxide, from which, finally, water is eliminated (Ballou et al., 2005). This catalytic process has diverse requirements that are difficult to be satisfied by a single catalytic site. Two general strategies have evolved to deal with this complex chemical problem (Ballou et al., 2005). First, in the case of single-component flavin monooxygenases, the enzyme undergoes significant protein and

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Aerobic Degradation of Aromatic Hydrocarbons

flavin dynamics during catalysis. The second approach uses two components to separate the catalytic tasks, an oxidoreductase to generate reduced flavin, and an oxygenase to receive the reduced flavin, react with O2 and hydroxylate the substrate (Ballou et al., 2005). Flavin monooxygenases have been classified according to sequence and structural data in six classes (van Berkel et al., 2006), with classes A, D and E being of special importance for aromatic hydrocarbon degradation. 2.1.3.1

Class A Single-Component Flavin Monooxygenases

Class A flavin monooxygenases are encoded by a single gene, contain a tightly bound FAD as cofactor, depend on NADH or NADPH as cofactor and are structurally composed of one dinucleotide binding domain to bind FAD. They are widely distributed in different bacterial taxa as hydroxylases in ortho- or para-position of aromatic compounds that already contain a hydroxyl group (van Berkel et al., 2006) (> Fig. 2). The 4-hydroxybenzoate 3-hydroxylase of P. fluorescens is one of the most thoroughly studied Class A enzymes (Entsch et al., 1987). This enzyme (encoded by the pobA gene) catalyzes the conversion of 4-hydroxybenzoate to protocatechuate. Typically PobA gene products show a narrow specificity and in addition to 4-hydroxybenzoate also hydroxylate 4-aminobenzoate to 4-amino-3-hydroxybenzoate (Entsch and van Berkel, 1995). The purification from several bacterial sources and the presence of pobA homologous genes in Actinomycetes, a, b, and gProteobacteria is indicative of a broad distribution of this enzyme. In a dendrogram of singlecomponent monooxygenases, PobA gene products are exclusively clustered without any additional sequences predicting a common evolutionary origin (> Fig. 2). Single component salicylate 1-hydroxylases catalyze the transformation of salicylate to catechol and was the first flavin monooxygenase characterized (Yamamoto et al., 1965). This enzyme has also been purified and characterized from many microorganisms showing a relatively broader specificity including chloro- and methylsalicylates as substrates (Camara et al., 2007; Lehrbach et al., 1984). Salicylate 1-hydroxylases clustered very close in the dendrogram with the exception of NahW, an isoenzyme found in P. stutzeri AN10 (Bosch et al., 1999) (> Fig. 2). Two distinct single component monooxygenases acting on 3-hydroxybenzoate have been described, 3-hydroxybenzoate 4-hydroxylase producing protocatechuate (Hiromoto et al., 2006) and 3-hydroxybenzoate 6-hydroxylase producing gentisate (Wang et al., 1987). The latter enzymes are closely related to salicylate 1-hydroxylases (> Fig. 2), and are, in genome sequencing projects often misleadingly annotated as salicylate hydroxylases. 3-Hydroxybenzoate 4-hydroxylases have so far only been reported in Comamonas strains and are only distantly related to other hydroxybenzoate hydroxylases. Another group of single component flavin monooxygenases belonging to class A, termed phenol hydroxylases, has been described from phenol degrading Pseudomonas strains, among them PheA from Pseudomonas sp. strain EST1001, which transforms phenol and 3-methylphenol (Nurk et al., 1991). This substrate specificity significantly differs from that described for a group of closely related enzymes termed 2,4-dichlorophenol hydroxylases (> Chapter 5, Vol. 2, Part 2). Various other closely related enzymes have confirmed or assumed activity as 2-hydroxybiphenyl 3-hydroxylase (HbpA of P. azelaica (Suske et al., 1997)), benzoquinol monooxygenases (chlorobenzoquinol monooxygenase of Pimelobacter simplex E3 (AY822041) and methylbenzoquinol monooxygenase of Burkholderia sp. NF100 (Tago et al., 2005)) or 2-hydroxyphenylpropionate 3-hydroxylase from Rhodococcus sp. V49 (Powell and Archer, 1998)) (see > Fig. 2). Enzymes of this group are commonly and misleadingly annotated as 2,4-dichlorophenol hydroxylases.

Aerobic Degradation of Aromatic Hydrocarbons

4

. Figure 2 Dendrogram showing the relatedness of single component flavoprotein monooxygenases. Reactions catalyzed by enzymes indicated are given to the exterior of the figure.

Not only above mentioned benzoquinol monooxygenases can catalyze hydroxylation of dihydroxylated aromatic benzoquinal since a resorcinol (1,3-dihydroxybenzene) monooxygenase forming hydroxybenzoquinol, only distantly related to all above mentioned enzymes has been identified in Corynebacterium glutamicum (Huang et al., 2006). Whereas a class A 2-hydroxyphenylpropionate 3-hydroxylase forming 2,3-dihydroxyphenylpropionate has thus far only been identified in Rhodococcus (Powell and Archer, 1998), class A 3-hydroxyphenylpropionate 2-hydroxylases forming the same product have been identified in Actinobacteria and Proteobacteria (Barnes et al., 1997; Ferra´ndez et al., 1997) and clustered together in dendrogram (> Fig. 2). Also 3-hydroxyphenylacetate is subject to monooxygenation by a 3-hydroxyphenylacetate 6-hydroxylase (> Fig. 2), giving rise to homogentisate (Arias-Barrau et al., 2005). The MhaA 3-hydroxyphenylacetate 6-hydroxylase of P. putida, however, necessitates the presence of

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MhaB, described as an essential coupling protein, for activity. Thus it constitutes a novel type of two-component hydroxylase (Arias-Barrau et al., 2005), distinct from the more classical two-component flavoprotein monooxygenases described below. 2.1.3.2

Two-Component Flavin Monooxygenases

During the past few years, several two-component aromatic hydroxylases consisting of an oxidoreductase and an oxygenase have been identified which have no structural or sequence similarities to the single-component enzymes and were classified as type D and E flavoprotein monooxygenases (Ballou et al., 2005). Class D flavoprotein monooxygenases (van Berkel et al., 2006) use FADH2 generated by the oxidoreductase as a coenzyme, show a structural resemblance with the acyl-CoA dehydrogenase fold, and comprise 4-hydroxyphenylacetate 3-hydroxylase (e.g., from E. coli, Acinetobacter baumanii or P. putida), phenol hydroxylase from Bacillus thermoglucosidasius (van Berkel et al., 2006), and also trichlorophenol monooxygenases (See > Chapter 5, Vol. 2, Part 2). Alternatively to 4-hydroxyphenylacetate 3-hydroxylase, a 1-hydroxylation of 4-hydroxyphenylacetate with homogentisate as reaction product has been described (Hareland et al., 1975), however, no information is available so far on the encoding gene nor on the detailed enzyme mechanism and thus on the class this enzyme belongs to. Styrene monooxygenase (StyA) has been identified from various Pseudomonas strains (Beltrametti et al., 1997), and was classified as Class E flavoprotein monooxygenase. An evolutionary link with the Class A flavoprotein monooxygenases was suggested (van Berkel et al., 2006). Styrene monooxygenases are highly enantioselective in oxidizing styrene and some of its derivatives to the respective epoxides.

2.2

Side Chain Processing

2.2.1

Oxidations of Methyl Groups in Methyl-Substituted Aromatic Hydrocarbons

Some bacteria catabolize methyl-substituted aromatic hydrocarbons such as toluene, xylenes (Assinder and Williams, 1990), p-cymene (DeFrank and Ribbons, 1976), and p-cresol (p-methylphenol) (Hopper and Taylor, 1977) by oxidizing the methyl group to the corresponding acids (> Fig. 3). The best known aromatic methyl-substituent oxidation pathway is that encoded by TOL plasmid, pWW0, in P. putida mt-2 (Assinder and Williams, 1990), involving oxidation of toluene, m-xylene, and p-xylene to benzoate, m-toluate, and p-toluate, respectively (> Fig. 3). The pathway is initiated by a monooxygenase, which catalyzes the oxidation of toluene (or m- or p-xylene) to benzyl alcohol. This monooxygenase is a two-component enzyme consisting of a XylA reductase subunit, which transfers electrons from NADH through FAD and a [2Fe-2S] center to the membrane-associated XylM hydroxylase subunit. There, one atom of activated molecular oxygen is inserted into the methyl group while the other oxygen atom is reduced to water. XylM shares significant amino acid identity (approx. 25%) with the integral-membrane non-heme diiron AlkB alkane hydroxylases. The conversion of benzyl alcohols to benzaldehydes is catalyzed by an NAD+-linked alcohol dehydrogenase. It seems that dehydrogenation is the major route for this transformation, however, the alcohol is presumably oxidized by the monooxygenase to an unstable gem-diol

Aerobic Degradation of Aromatic Hydrocarbons

4

. Figure 3 Peripheral reactions involved in oxidation of methyl groups in methyl-substituted aromatic hydrocarbons. Reaction intermediates are shown in brackets.

intermediate which is recognizable as the hydrate of the corresponding benzaldehyde (Harayama et al., 1986). The aldehyde formed is then oxidized to benzoate (or m-toluate or p-toluate), by an NAD+-linked aldehyde dehydrogenase. Three enzymes -a two-component monooxygenase, an alcohol dehydrogenase, and an aldehyde dehydrogenase- have also been involved in the catabolism of p-cymene through p-cumate (p-isopropylbenzoate) (DeFrank and Ribbons, 1976; Eaton, 1997) (> Fig. 3). However, only the monooxygenase and aldehyde dehydrogenase of the p-cymene catabolic pathway are related to the analogous enzymes of the TOL plasmid (Eaton, 1997). The metabolism of p-cresol, as studied in P. putida NCIB 9866 (Hopper, 1976), is initiated by a periplasmatic p-cresol methylhydroxylase (PCMH) to p-hydroxybenzaldehyde with the transient formation of p-hydroxybenzyl alcohol (Keat and Hopper, 1978). This enzyme, a flavocytochrome, consists of an a subunit containing the active site and a FAD covalently linked to tyrosine, and a c-type cytochrome containing b subunit (McIntire et al., 1985). The product of PCMH, p-hydroxybenzaldehyde, is oxidized by a dehydrogenase to p-hydroxybenzoate (Cronin et al., 1999). 4-Ethylphenol methylenehydroxylase from P. putida JD1 is related to

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PCMH (Reeve et al., 1989) (> Fig. 3). Catabolism of 4-ethylphenol in strain JD1 proceeds by hydroxylation to give 1-(40 -hydroxyphenyl)ethanol, followed by dehydrogenation to the ketone, 4-hydroxyacetophenone (Darby et al., 1987) (> Fig. 3).

2.2.2

O-Demethylations

Methoxylated aromatic hydrocarbons such as vanillate or syringate are important intermediate metabolites from lignin. Some demethylating systems have been described in aerobic bacteria to deal with such metabolites. Vanillate O-demethylase belongs to the Rieske nonheme iron oxygenases (see > Fig. 1). It is a two-component oxygenase consisting of a reductase (VanB) and an oxygenase (VanA). The oxygenase is composed only of a-subunits and shares similarity with phthalate 4,5-dioxygenases (see 2.1.1. and > Fig. 1). This type of demethylase is involved in vanillate degradation by most vanillate-degrading aerobic bacteria, such as Pseudomonas and Acinetobacter strains (Buswell and Ribbons, 1988). Vanillate demethylases have a wide substrate specificity and vanillate analogs that are transformed share the common property of a methoxy- or methyl group in meta-position to the carboxyl group (such as m-anisate, 3,4-dimethoxybenzoate or m-toluate) (Morawski et al., 2000). The enzyme is not only able to demethylate methoxy groups but also to monohydroxylate methyl groups in the meta-position. Another type of demethylase is the tetrahydrofolate (THF) dependent demethylases, which have mainly been reported in anaerobic bacteria. However, THF dependent syringate and vanillate O-demethylases have also been reported in Sphingomonas paucimobilis SYK-6 (Abe et al., 2005; Masai et al., 2004) (> Fig. 4). The deduced amino acid sequence of DesA syringate O-demethylase shows similarity to the THF-dependent aminomethyltransferase of E. coli involved in glycine cleavage. DesA converts syringate to 3-O-methylgallate only in the presence of THF, with the concomitant formation of 5-methyl-THF. Vanillate and 3-O-methylgallate are also used as substrates for DesA, however with poor activity (Masai et al., 2004). More recently, a second THF dependent O-demethylase termed LigM, showing 49% of amino acid sequence identity with DesA, was discovered in S. paucimobilis SYK-6. In the presence of THF, LigM converts vanillate and 3-O-methylgallate into protocatechuate and gallate, respectively (> Fig. 4), whereas syringate was not transformed (Abe et al., 2005). Cytochrome P450 O-demethylase systems have been described for the demethylation of veratrole and guaiacol to catechol in Streptomyces setonii and Moraxella sp. respectively (Sauret-Ignazi et al., 1988; Sutherland, 1986), and for demethylation of 2-ethoxyphenol and 4-methoxybenzoate in Rhodococcus rhodochrous (Karlson et al., 1993), however, identification of the cytochrome P450 genes is lacking.

2.2.3

Aromatic Acid Decarboxylations

Decarboxylations are required for degradation of phthalate, 5-carboxyvanillate and 2,6dihydroxybenzoate, among other aromatic acids. Decarboxylases involved in the elimination of the carboxyl group from the aromatic nucleus in these pathways have been reported as nonoxidative (reductive) decarboxylases that do not require the external addition of any cofactor for its activity.

Aerobic Degradation of Aromatic Hydrocarbons

4

. Figure 4 Metabolism of methoxylated aromatic hydrocarbons by Sphingomonas paucimobilis SYK-6. Enzymes catalyzing a given reaction only at slow rate are shown in italics.

Phthalate is metabolized by two different dioxygenase-initiated pathways (see 2.1.1) either via 4,5-dihydroxyphthalate (in Proteobacteria) or 3,4-dihydroxyphthalate (in Actinobacteria) (> Fig. 1). Both dihydroxyphthalate isomers are non-oxidatively decarboxylated to protocatechuate. 4,5-dihydroxyphthalate decarboxylases have been purified from P. fluorescens PHK (Pujar and Ribbons, 1985) and C. testosteroni NH1000 (Nakazawa and Hayashi, 1978) and show a narrow specificity with only 4,5-dihydroxyphthalate and 4-hydroxyphthalate as substrates (Nakazawa and Hayashi, 1978). 4,5-Dihydroxyphthalate decarboxylases described thus far share >78% of sequence identity. 3,4-dihydroxyphthalate 2-decarboxylase from A. keyseri 12B is unrelated to the 4,5-dihydroxyphthalate decarboxylases and its deduced amino acid

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Aerobic Degradation of Aromatic Hydrocarbons

sequence most closely resembles that of aldolases which catalyze the cleavage of fuculose 1-phosphate (Eaton, 2001). Similar 3,4-dihydroxyphthalate 2-decarboxylases (58–69% identity) were observed in other phthalate-degrading Actinobacteria. In S. paucimobilis SYK-6, 5-carboxyvanillate is transformed to vanillate by non-oxidative 5-carboxyvanillate decarboxylases (> Fig. 4). Two such decarboxylases, LigW and LigW2, have been identified, which share 37% amino acid sequence identity (Peng et al., 2002, 2005) but exhibit no homology with members of previously described non-oxidative decarboxylase families. It was recently proposed that they can be classified into a new family of non-oxidative aromatic acid decarboxylases, together with 2,6-dihydroxybenzoate decarboxylase of Rhizobium radiobacter (Yoshida et al., 2004), which catalyzes the reversible decarboxylation of 2,6- and 2,3-dihydroxybenzoate to resorcinol and catechol, respectively.

2.2.4

CoA-Dependent Peripheral Pathways for Phenylpropenoid Compounds

Phenylpropenoid compounds constitute a common carbon source for plant-associated microorganisms since they are structural components of plant polymers, such as lignin and suberin. Among phenylpropenoid compounds, the largest group is hydroxycinnamates derivatives (i.e., ferulate, coumarate, caffeate and others). In A. baylyi ADP1, a chlorogenate esterase hydrolyzes the ester bond of chlorogenate, an abundant hydroxycinnamic compound, to produce quinate and caffeate (Smith et al., 2003). In this bacterium, like in most other bacteria characterized in the respect, hydroxycinnamate catabolism follows a CoA-dependent non b-oxidative route (Overhage et al., 1999). p-Coumarate, caffeate and ferulate are transformed to 4hydroxybenzoate, protocatechuate and vanillate, respectively, through the action of enzymes with relatively broad substrate specificity, encoded in A. baylyi by the hcaABC genes (> Fig. 5). Hydroxycinnamates are initially activated to the corresponding CoA esters by hydroxycinnamoyl-CoA synthase (HcaC) (also often termed feruloyl-CoA synthase), an ATP dependent (AMP forming) CoA ligase (> Fig. 5). A bifunctional enoyl-CoA hydratase/aldolase (HcaA) catalyzes the hydratation but also acts as a lyase cleaving the hydrated derivatives of feruloyl-CoA, caffeoyl-CoA and p-coumaroyl-CoA to acetyl-CoA and vanillin, 3,4-dihydroxybenzaldehyde and 4-hydroxybenzaldehyde, respectively (Gasson et al., 1998). The aldehydes are oxidized by aldehyde dehydrogenases (HcaB) to vanillate, protocatechuate, and 4-hydroxybenzoate, respectively (> Fig. 5). 4-Hydroxybenzoate and vanillate are transformed to protocatechuate by 4-hydroxybenzoate 3-hydroxylase or vanillate O-demethylase, respectively. Three alternative modes of ferulate degradation are additionally discussed in the literature, a non-oxidative decarboxylation, discovered mainly in fungi and yeast, side-chain reduction, typical for the anaerobic degradation of ferulate, and a CoA-independent deacetylation (Priefert et al., 2001). However, even in Delftia acidovorans, which had been proposed to carry out such a CoA-independent deacetylation, a CoA-dependent pathway was observed as the major route for ferulate degradation (Plaggenborg et al., 2001). Phenylpropanoid compounds, i.e., saturated derivatives of hydroxycinnamates, such as 4-hydroxyphenylpropionate and 3,4-dihydroxyphenylpropionate are assumed to be catabolized by A. baylyi ADP1, using a FAD-dependent acyl-CoA dehydrogenase (HcaD) which dehydrogenates the saturated propionyl-CoA side chain of the hydroxyphenylpropanoyl thioesters produced by HcaC to form hydroxycinnamoyl-CoA thioesters that can be channeled to protocatechuate (Smith et al., 2003).

Aerobic Degradation of Aromatic Hydrocarbons

. Figure 5 Peripheral reactions in the bacterial degradation of ferulate, caffeate, p-coumarate, phenylalanine and tryptophan.

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Aerobic Degradation of Aromatic Hydrocarbons

Peripheral Reactions in the Degradation of Aromatic Amino Acids

In eukaryotes, the metabolism of the aromatic amino acids phenylalanine, tyrosine and tryptophan is initiated by tetrahydropterin dependent monooxygenases (Fitzpatrick, 2003), where tetrahydropterin serves as electron source to reduce the second atom of oxygen to the level of water. Also in bacteria, phenylalanine is transformed by a pterin-dependent phenylalanine hydroxylase into tyrosine (Nakata et al., 1979) (> Fig. 5). A tyrosine aminotransferase catalyzes the conversion of tyrosine into 4-hydroxyphenylpyruvate (Arias-Barrau et al., 2004; Gu et al., 1998), which is further transformed by a 4-hydroxyphenylpyruvate dioxygenase (HPPD) (Fitzpatrick, 2003). HPPD is an Fe2+-dependent, non-heme oxygenase that catalyzes the conversion of 4-hydroxyphenylpyruvate to homogentisate (> Fig. 5). This reaction involves decarboxylation, substituent migration and aromatic oxygenation in a single catalytic cycle. This enzyme is a member of the a-keto acid dependent oxygenases that typically require an a-keto acid (almost exclusively a-ketoglutarate) and molecular oxygen to either oxygenate or oxidize a third molecule. As an exception in this class of enzymes HPPD has only two substrates, does not use a-ketoglutarate, and incorporates both atoms of oxygen into the aromatic product, homogentisate (Moran, 2005). Indications were also given that phenylalanine, in an alternative pathway can be metabolized via 3,4-dihydroxyphenylalanine and protocatechuate (Ranjith et al., 2007). In various bacteria, tryptophan is subject to non-oxidative degradation by a pyridoxal phosphate-dependent tryptophan indole-lyase (tryptophanase) yielding indole, pyruvate and ammonium (Vederas et al., 1978). In some bacteria, the oxidative degradation of exogenous tryptophan via anthranilate has been suggested (Kurnasov et al., 2003a), but details are still scarce. Experimental verification was achieved by functional expression of a Cupriavidus metallidurans putative kynBAU operon. Tryptophan is converted by a heme-containing specific tryptophan 2,3-dioxygenase (KynA) to N-formylkynurenine, from which the formyl group is removed by kynurenine formamidase (KynB) to kynurenine, and kynureninase (KynU) catalyzes the cleavage to anthranilate and alanine (Kurnasov et al., 2003a) (> Fig. 5). However, further genome analyses revealed the presence of gene clusters encoding kynurenine monooxygenase together with kynureninase and a 3-hydroxyanthranilate 3,4dioxygenase (see 2.3.5.4) (Kurnasov et al., 2003b) indicating kynurenine to be monooxygenated to 3-hydroxykynurenine followed by cleavage to alanine and 3-hydroxyanthranilate by kynureninase (> Fig. 5), which accepts both kynurenine and 3-hydroxykynurenine as substrates (Kurnasov et al., 2003b). Thus, tryptophan can be degraded in bacteria via anthranilate, but also via 3-hydroxyanthranilate. How these different pathways contribute to tryptophan degradation in different taxa remains to be established.

2.3

Central Reactions

2.3.1

Intradiol Ring-Cleavage Pathways

2.3.1.1

The 3-Oxoadipate Pathway

The 3-oxoadipate pathway is widely distributed among soil bacteria and plays a central role in the degradation of naturally occurring aromatic hydrocarbons (Harwood and Parales, 1996) such as vanillin, p-coumarate, caffeate, mandelate or tryptophan. Two branches of the

Aerobic Degradation of Aromatic Hydrocarbons

4

3-oxoadipate pathway can be differentiated, the catechol branch and the protocatechuate branch (> Fig. 6). In the catechol branch, the metabolism of catechol is initiated by ortho-cleavage catalyzed by catechol-1,2-dioxygenases resulting in the formation of cis,cis-muconate, which is subsequently transformed by a muconate cycloisomerase to muconolactone. Muconolactone isomerase shifts the double bond to form 3-oxoadipate-enol-lactone (enol-lactone), the first common intermediate of the catechol and protocatechuate branch (> Fig. 6). In the protocatechuate branch, protocatechuate is subject to ortho-cleavage by protocatechuate 3,4-dioxygenases. Like catechol 1,2-dioxygenases, protocatechuate 3,4-dioxygenases are non-heme Fe3+ containing dioxygenases (Fujisawa and Hayaishi, 1968). However, in contrast to catechol 1,2-dioxygenases, which are composed of only one type of subunits, protocatechuate 3,4-dioxygenases are composed of two different subunits which, however, share substantial amino acid identity (Yoshida et al., 1976). Even though cycloisomerization is an important step in both branches of the 3-oxoadipate pathway, the enzymes catalyzing the respective reactions are different. Sequence analyses and kinetic studies showed that carboxymuconate cycloisomerases of the protocatechuate branch belong to the fumarase class II family (Williams et al., 1992), a group of enzymes catalyzing 1,2-addition–elimination reactions including aspartase and arginosuccinate lyase. They do not require any metal cofactors for catalytic activity and catalyze a syn-1,2-addition-elimination with 4-carboxy-(S)-muconolactone as product. In contrast, the Mn2+ requiring muconate cycloisomerases belong to the enolase superfamily and catalyze an anti-1,2-addition-elimination with (R)-muconolactone as product (Babbitt et al., 1996). In the protocatechuate branch, the 4-carboxymuconolactone produced is transformed to enol-lactone by carboxymuconolactone decarboxylase. From the biochemical point of view, both branches merge at the stage of the enol-lactone (> Fig. 6), however, a remarkable diversity of the 3-oxoadipate pathway in terms of gene organization, type of inducers and regulation mechanism has been observed (Harwood and Parales, 1996), such that the pathways genetically converge at different points in different bacteria, or they never converge as in A. baylyi, which contains two independent set of genes encoding isofunctional enzymes for the last three steps of the pathway. The pcaC and pcaD genes, encoding 4-carboxymuconolactone decarboxylase and 3-oxoadipate enol-lactone hydrolase, respectively, catalyzing successive reactions, are, in some cases, fused in a unique pcaL gene. Sequence analysis of pcaL genes reveals that the N-terminal two thirds of the protein are homologous to the enol-lactone hydrolases, whereas the C-terminal third is homologous to the decarboxylases (Eulberg et al., 1998). As these gene fusions are present in distantly related bacteria a biochemical advantage of these fused gene products is possible (Pe´rez-Pantoja et al., 2008). Enol-lactone is hydrolyzed by enol-lactone hydrolases which are assumed to use a Ser-His-Asp catalytic triade (Schlo¨mann, 1994). 3-Oxoadipate in turn is transformed by 3-oxoadipate:succinyl-CoA transferase and 3-oxoadipyl-CoA thiolase to Krebs cycle intermediates (Gobel et al., 2002). 2.3.1.2

Metabolism of Methylaromatics via Intradiol Cleavage

Usually, the 3-oxoadipate pathway is not suited for the degradation of methylaromatics because methylsubstituted muconolactones, formed by the action of catechol 1,2-dioxygenase and muconate cycloisomerase, accumulate as dead-end products (Catelani et al., 1971). In the case of transformation of 4-methylcatechol, 4-methylmuconolactone (4-ML) is formed (> Fig. 6), which cannot be processed by enzymes of the 3-oxoadipate pathway as no proton is available to be abstracted by the muconolactone isomerase (Pieper et al., 1985). However, in

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. Figure 6 Dendrogram showing the relatedness of intradiol dioxygenases (catechol 1,2-dioxygenases, protocatechuate 3,4-dioxygenases and hydroxybenzoquinol 1,2-dioxygenases). Reactions catalyzed by enzymes indicated are given to the exterior of the figure, together with subsequent reactions channeling the ring-cleavage products into the Krebs cycle. Among catechol 1,2-dioxygenases, two different lineages can be differentiated, observed in Proteobacteria and Actinobacteria, respectively (Eulberg et al., 1997). Reactions of catechol pathway enzymes with 4-methylcatechol resulting in the formation of 4-methylmuconolactone are indicated to the left of the figure.

Aerobic Degradation of Aromatic Hydrocarbons

4

C. necator JMP134 and R. rhodochrous N75, a 4-methylmuconolactone methylisomerase capable of converting 4-ML to 3-methylmuconolactone (3-ML) was described which compensates for the initial ‘‘incorrect’’ cycloisomerization of 3-methylmuconate (Bruce et al., 1989; Pieper et al., 1990). In C. necator JMP134, 3-ML is further metabolized by a methylmuconolactone isomerase and via 4-methyl-3-oxoadipate, and hence, probably, by analogous reactions to those of the classical 3-oxoadipate pathway (Prucha et al., 1997). However, further metabolism of 3-ML is still poorly understood, and in R. rhodochrous N75 hydrolysis of the lactone ring obviously occurs from 3-methylmuconolactone-CoA (Cha et al., 1998). 2.3.1.3

Metabolism of 1,2,4-Trihydroxybenzene

Hydroxybenzoquinol (1,2,4-trihydroxybenzene) is the central intermediate in the degradation of a variety of aromatic hydrocarbons such as resorcinol (Huang et al., 2006), 4-aminophenol (which is assumed to be degraded via 1,4-benzenediol, (Takenaka et al., 2003)) or 4-hydroxysalicylate (Armengaud et al., 1999) including a variety of particularly recalcitrant polychloro- and nitroaromatic pollutants (> Chapter 5, Vol. 2, Part 2). Hydroxybenzoquinol 1,2-dioxygenase is the key enzyme of hydroxybenzoquinol metabolism and catalyzes the intradiol cleavage to form 3-hydroxy-cis,cis-muconate and its tautomer, maleylacetate (> Fig. 6). Hydroxybenzoquinol 1,2-dioxygenases have been purified and characterized from a variety of microorganisms (Takenaka et al., 2003), and also crystallized (Ferraroni et al., 2005). Hydroxybenzoquinol 1,2-dioxygenases are usually highly specific for hydroxybenzoquinol and do not, or relatively slowly, convert catechol. In accordance, in a dendrogram of intradiol dioxygenases, hydroxybenzoquinol and catechol 1,2-dioxygenases clustered in separated branches (> Fig. 6). The next enzyme of the hydroxyquinol pathway, maleylacetate reductase, performs the reduction of the carbon-carbon double bond to channel maleylacetate into the 3-oxoadipate pathway. Maleylacetate reductases have previously been described as important key enzymes of chloroaromatics but also nitroaromatics degradation (See > Chapter 5, Vol. 2, Part 2).

2.3.2

Catechol Meta-Cleavage Pathways

The extradiol ring-cleavage of catechol and methylsubstituted catechols is typically catalyzed by type I extradiol dioxygenases (catechol 2,3-dioxygenases, C23O) which belong to the vicinal oxygen chelate family enzymes (Gerlt and Babbitt, 2001). Type I extradiol dioxygenases are also involved in the degradation of biphenyl (2,3-dihydroxybiphenyl 1,2-dioxygenases, See > Chapter 26, Vol. 2, Part 5), or naphthalene (See > Chapter 24, Vol. 2, Part 5). All these extradiol dioxygenases, use non-heme Fe2+ for cleavage (Eltis and Bolin, 1996; Harayama and Rekik, 1989). However, Mn2+ dependent extradiol dioxygenases with high sequence similarity to the Fe2+ dependent enzymes have also been reported (Hatta et al., 2003). Eltis and Bolin (Eltis and Bolin, 1996) analyzed in detail the phylogenetic relationships among type I extradiol dioxygenases and described them as a superfamily which can be divided into different families and subfamilies. Families I.2 and I.3 consist of two-domain iron-containing enzymes that show preferences for monocyclic and bicyclic substrates, respectively, whereas family I.1 comprises the small single domain enzymes identified in R. globerulus P6 and Sphingomonas sp. strain BN6. In the last years, the description of new members of this family increased significantly (Vaillancourt et al., 2006).

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Extradiol dioxygenases can be subject to a rapid oxidation of the active site ferrous iron into its ferric form with concomitant loss of activity (Vaillancourt et al., 2002), specifically during turnover of certain substrates such as 4-methylcatechol. Small auxiliary ferredoxin proteins, whose genes are frequently encoded adjacently to the C23O genes, have been reported to have a reactivating function through reduction of the iron atom in the active site of the enzyme (Hugo et al., 1998, See > Chapter 19, Vol. 2, Part 4). Two branches of the meta-cleavage pathway for catechols have been described, the hydrolytic and the oxalocrotonate branch, which are often encoded in one single gene cluster (Harayama et al., 1987). In the oxalocrotonate branch, 2-hydroxymuconic semialdehyde is oxidized to 2-hydroxymuconate by 2-hydroxymuconic semialdehyde dehydrogenase, followed by isomerization to oxalocrotonate through the action of oxalocrotonate isomerase and decarboxylation by oxalocrotonate decarboxylase to 2-hydroxypent-2,4-dienoate, the common intermediate of the hydrolytic and the 4-oxoalocrotonate branch (> Fig. 7). Both 2-hydroxymuconic semialdehyde (from catechol) and 5-methyl-2-hydroxymuconic semialdehyde (from 4-methylcatechol) are preferentially degraded via the oxalocrotonate branch (Harayama et al., 1987). Since 2-hydroxy-6-oxo-2,4-heptadienoate, the ring-cleavage product of 3-methylcatechol is a ketone, rather than an aldehyde, it cannot be further oxidized by the 2-hydroxymuconic semialdehyde dehydrogenase and is exclusively metabolized via the hydrolytic route (Powlowski and Shingler, 1994). Hydrolysis of 2-hydroxy-6-oxo-2,4-heptadienoate by 2-hydroxymuconic semialdehyde hydrolase gives rise to 2-hydroxypent-2,4dienoate and acetate (> Fig. 7). The final steps of the catechol meta-cleavage pathway are catalyzed by 2-hydroxypent-2,4-dienoate hydratase (to give 4-hydroxy-2-oxovalerate), 4-hydroxy-2-oxovalerate aldolase (to give acetaldehyde and pyruvate) and acetaldehyde dehydrogenase (decycling) that converts acetaldehyde to acetyl-CoA.

2.3.3

Protocatechuate Meta-Cleavage Pathways

Like for catechol, protocatechuate can be metabolized via intradiol or extradiol cleavage pathways. As protocatechuate has an asymmetric structure, meta-cleavage can occur in the 2,3- but also in the 4,5-position. Most microorganisms seem to perform a 4,5-cleavage (Ono et al., 1970). A protocatechuate 2,3-dioxygenase has been described from Bacillus macerans (Wolgel et al., 1993), however, no further information on this pathway is yet available. Protocatechuate 4,5-cleavage is catalyzed by heteromultimeric protocatechuate 4,5-dioxygenases, with the two subunits being unrelated (Sugimoto et al., 1999). The active site comprises, like in C23O, a Fe2+ ion located in the b-subunit (Sugimoto et al., 1999). However, protocatechuate 4,5-dioxygenase is unrelated to above described C23Os and belongs to the type II or LigB superfamily of extradiol dioxygenases (Vaillancourt et al., 2006) (> Fig. 8). The protocatechuate 4,5-dioxygenolytic ring-cleavage product 4-carboxy-2-hydroxymuconate-6-semialdehyde is non enzymatically converted to an intramolecular hemiacetal form and then oxidized by a 4-carboxy-2-hydroxymuconic semialdehyde dehydrogenase (> Fig. 7) which, like cis-4,5-dihydro-4,5-dihydroxyphthalate dehydrogenase (see 2.1.1), belongs to the GFO/IDH/MOCA family (Chang and Zylstra, 1998). The resulting intermediate, 2-pyrone4,6-dicarboxylate, is hydrolyzed by 2-pyrone-4,6-dicarboxylate hydrolase (Maruyama, 1983) to yield the keto form and enol form (4-carboxy-2-hydroxymuconate) of 4-oxalomesaconate, which are in equilibrium. 2-Pyrone-4,6-dicarboxylate hydrolase was postulated to contain a catalytically active cysteine as part of a catalytic triad (Masai et al., 1999b), however, further

. Figure 7 Extradiol ring-cleavage pathways involved in the degradation of catechol, 3-methylcatechol, protocatechuate, homoprotocatechuate, 2,3dihydroxyphenylpropionate, 2,3-dihydroxycinnamate, 2-aminophenol, 3-hydroxyanthranilate and 4-amino-3-hydroxybenzoate. Unstable intermediates are shown in brackets. In case of catechol only metabolism via the oxalocrotonate branch is indicated.

Aerobic Degradation of Aromatic Hydrocarbons

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. Figure 8 Dendrogram showing the relatedness of type II (LigB) extradiol dioxygenases. The physiological function of BphC6 2,3-dihydroxybiphenyl 1,2dioxygenase from R. rhodochrous K37 remains to be established, but is possibly involved in fluorene degradation. (Taguchi et al., 2004)

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analysis on related proteins indicates that this hydrolase might be a metal-dependent hydrolase (Halak et al., 2007). The recent elucidation of the crystal structure might shed some light on the enzyme mechanism. 4-Oxalomesaconate is converted to 4-carboxy-4-hydroxy2-oxoadipate by 4-oxalomesaconate hydratase (Hara et al., 2000). Finally, 4-carboxy-4hydroxy-2-oxoadipate is cleaved by 4-carboxy-4-hydroxy-2-oxoadipate aldolase to produce pyruvate and oxaloacetate (> Fig. 7).

2.3.4

Further Meta-Cleavage Routes Involving Type I or Type II Extradiol Dioxygenases

2.3.4.1

The Homoprotocatechuate Pathway

Homoprotocatechuate (3,4-dihydroxyphenylacetate) is a central intermediate in the degradation of 4-hydroxyphenylacetate and the aromatic amines tyramine and dopamine. So far, degradation of homoprotocatechuate has been exclusively described by extradiol cleavage through homoprotocatechuate 2,3-dioxygenases. Proteobacterial homoprotocatechuate 2,3-dioxygenases as the one described from E. coli C (Roper and Cooper, 1990) belong to the type II or LigB superfamily of extradiol dioxygenases (> Fig. 8). In contrast, actinobacterial homoprotocatechuate 2,3-dioxygenases like the Fe2+ dependent enzyme from Brevibacterium fuscum or the Mn2+ dependent enzyme from Arthrobacter globiformis belong to the type I extradiol dioxygenases (Vetting et al., 2004). Independent of the type of reaction, 5-carboxymethyl-2-hydroxymuconic semialdehyde is the reaction product (> Fig. 7). In E. coli, the further metabolism follows a dehydrogenative route with dehydrogenation to the acid by 5-carboxymethyl-2-hydroxymuconic semialdehyde dehydrogenase, which exhibits significant sequence identity (40%) to the respective 2-hydroxymuconic semialdehyde dehydrogenases involved in catechol degradation (Diaz et al., 2001). Isomerization of 5-carboxymethyl-2-hydroxymuconate to 5-oxopent-3-ene-1,2,5-tricarboxylic acid is catalyzed by an isomerase in a reaction similar to that performed by 4-oxalocrotonate tautomerase (see 2.3.2), however, the two enzymes do not have any apparent sequence similarity (Diaz et al., 2001). A bifunctional decarboxylase/isomerase catalyzes the magnesium-dependent decarboxylation of 5-oxopent-3-ene-1,2,5-tricarboxylate to 2-oxo-hept-3-ene-1,7-dioate. This reaction is followed by a hydratase giving rise to 2,4-dihydroxyhept-2-ene-1,7-dioate and an aldolase forming pyruvate and succinic semialdehyde (> Fig. 7). 2.3.4.2

The 2,3-Dihydroxyphenylpropionate Pathway

Extradiol ring-cleavage is also involved in the metabolism of 3-hydroxyphenylpropionate and 3-hydroxycinnamate via 2,3-dihydroxyphenylpropionate or 2,3-dihydroxycinnamate, respectively. Like proteobacterial 3,4-dihydroxyphenylacetate 2,3-dioxygenases, proteobacterial 2,3-dihydroxyphenylpropionate 1,2-dioxygenases belong to the LigB superfamily of extradiol dioxygenases (Diaz et al., 2001) (> Fig. 8). Similarly, actinobacterial 2,3-dihydroxyphenylpropionate 1,2-dioxygenases also belongs to this superfamily (Barnes et al., 1997). 2,3-dihydroxyphenylpropionate 1,2-dioxygenases show a broad specificity with 2,3-dihydroxycinnamate as a good substrate (Spence et al., 1996). Also catechol and methylcatechols are usually accepted as substrates (Barnes et al., 1997; Diaz et al., 2001). The ring-cleavage product 2-hydroxy-6-ketonona-2,4-diene-1,9-dioate is further degraded through a hydrolytic route generating succinate and 2-hydroxypent-2,4-dienoate (> Fig. 7). The respective hydrolases show some substrate selectivity for the carboxylate of the side chain

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with only slow turnover of the ring fission products of 3-methylcatechol or catechol (Diaz et al., 2001). However, the ring fission product of 2,3-dihydroxycinnamate was a fairly efficient substrate, generating fumarate and 2-hydroxypent-2,4-dienoate (Barnes et al., 1997; Lam and Bugg, 1997) (> Fig. 7). Significant sequence similarity has been detected between 2-hydroxy6-ketonon-2,4-diene-1,9-dioate hydrolases and other C-C bond hydrolases cleaving vinylogous 1,5-diketones such as those involved in the degradation of 2,3-dihydroxybiphenyl or catechol (see 2.3.2). 2-Hydroxy-6-ketonon-2,4-diene-1,9-dioate hydrolases of Proteobacteria seem to be most closely related to hydrolases involved in 2,3-dihydroxybiphenyl degradation, and share only approx. 30% sequence identity with 2-hydroxy-6-ketonon-2,4-diene-1,9dioate hydrolases of Actinobacteria. Further metabolism of 2-hydroxypent-2,4-dienoate occurs as described above (see 2.3.2). 2.3.4.3

Degradation of Gallate

Only poor information is available so far on the metabolism of gallate (3,4,5-trihydroxybenzoate). In S. paucimobilis SYK-6, it has been described that syringate is metabolized via 3-O-methylgallate (Masai et al., 1999a). The protocatechuate 4,5-dioxygenase of this strain was reported to catalyze ring-cleavage also of 3-O-methylgallate with the direct formation of 2-pyrone-4,6-dicarboxylate, a metabolite of protocatechuate degradation (see 2.3.3) (> Fig. 4). However, further analysis revealed the presence of a novel extradiol dioxygenase of the LigB family termed DesZ to be responsible for 3-O-methylgallate metabolism (Kasai et al., 2004). This enzyme also transforms gallate, but is practically inactive with protocatechuate (> Fig. 4). However, 3-O-methylgallate formed from syringate can be subject to an initial demethylation in strain SYK-6, and a third extradiol dioxygenase of the LigB family, termed DesB, could be identified as being highly specific for cleavage of gallate, and being inactive with either 3-O-methylgallate or protocatechuate (Kasai et al., 2005). Oxalomesaconate was identified as ring-cleavage product (> Fig 4). A similar gallate dioxygenase was also identified in P. putida KT2440 (Nogales et al., 2005) (> Fig. 8). A more detailed analysis of the primary structure of gallate dioxygenases revealed that the N-terminal regions showed a significant amino acid sequence identity with the b-subunit of protocatechuate 4,5-dioxygenases, whereas the C-terminal region has similarity to the corresponding small a-subunit (Nogales et al., 2005). It was therefore suggested that gallate dioxygenases are two-domain proteins that have evolved from the fusion of large and small subunits of protocatechuate 4,5-dioxygenases. 2.3.4.4

Degradation of 2-aminophenol

Usually, extradiol dioxygenases necessitate the presence of two neighbored hydroxyl-substituents on the substrate. However, analysis of the metabolism of 2-aminophenol revealed that this substrate can be directly cleaved by extradiol dioxygenases of the LigB superfamily, termed 2-aminophenol 1,6-dioxygenases (Takenaka et al., 1997) (> Fig. 8). These enzymes are composed of two subunits, which share sequence similarity (> Fig. 8) but it appears that only the b-subunit contains an active site. Catechol is only a poor substrate for the enzyme (Takenaka et al., 1997). The further metabolism of the formed 2-aminomuconic semialdehyde occurs in analogy to the metabolism of 2-hydroxymuconic semialdehyde produced during catechol extradiol cleavage (> Fig. 7). 2-aminomuconic semialdehyde dehydrogenases share significant sequence similarity (up to 60%) with 2-hydroxymuconic semialdehyde dehydrogenases and the enzyme of P. pseudoalcaligenes JS45 was shown to be capable to transform 2-hydroxymuconic semialdehyde (He et al., 1998). Aminomuconate is hydrolyzed by aminomuconate deaminase to 4-oxalocrotonate in strain JS45, which indicates that

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deamination is carried out via an imine intermediate (He and Spain, 1998). Aminomuconate deaminase was also observed in the degradation of 2-aminophenol by other Pseudomonas strains (Takenaka et al., 2000). Further degradation of 4-oxalocrotonate proceeds through reactions as described above (see 2.3.2) (> Fig. 7). A different aminomuconate deaminating activity was recently observed in Comamonas strain CNB-1 (Liu et al., 2007) and it was suggested that 2-hydroxymuconate rather than oxalocrotonate is the deamination product. 2.3.4.5

Extradiol Cleavage of Benzoquinol

Pathways that involve the cleavage of benzoquinol have been suggested to be involved in the degradation of 4-hydroxyphenoxyacetate (Crawford, 1978), 4-ethylphenol and 4-hydroxyacetophenone (Darby et al., 1987), but also of chloro- or nitroaromatics (See > Chapter 5, Vol. 2, Part 2). Chlorobenzoquinol (and also benzoquinol) produced during the catabolism of g-HCH (hexachlorocyclohexane) in S. japonicum UT26 is subject to direct ring cleavage by LinE extradiol dioxygenase, a type I extradiol dioxygenase (See > Chapter 5, Vol. 2, Part 2). In contrast, benzoquinol 1,2-dioxygenase from the 4-hydroxyacetophenone-degrading P. fluorescens ACB has been shown to be an a2b2 heterotetramer where the a- and b-subunits displayed no significant sequence identity with known dioxygenases. The enzyme is thus the prototype of a novel class of Fe2+-dependent dioxygenases (Moonen et al., 2008b). The enzyme not only cleaves benzoquinol to form 4-hydroxymuconic semialdehyde but also a wide range of substituted benzoquinols to the corresponding 4-hydroxymuconic semialdehyde derivatives. In P. fluorescens ACB, the subsequent conversion of 4-hydroxymuconic semialdehyde to maleylacetate is accomplished by a 4-hydroxymuconic semialdehyde dehydrogenase, which exhibits moderate sequence identity (37–43%) to the respective 2-hydroxymuconic semialdehyde dehydrogenases involved in meta-cleavage pathways of catechol (Moonen et al., 2008a). Maleylacetate is transformed to 3-oxoadipate by maleylacetate reductase to be channeled to Krebs cycle intermediates (See > Chapter 5, Vol. 2, Part 2).

2.3.5

Pathways Involving Extradiol Ring-Cleavage by Enzymes of the Cupin Superfamily

Proteins of the cupin superfamily share a common architecture and the term cupin (from the latin term ‘‘cupa,’’ for a small barrel or cask) has been given to a beta barrel structural domain (Dunwell et al., 2000). Members of this superfamily share two histidine containing sequence motifs that identify the binding site of the metal. Various extradiol dioxygenases of aromatic degradation pathways (termed type III) have been described to belong to this superfamily. It should be noted that even though belonging to different families, all three types of extradiol dioxygenases share similar active sites and all type I, type II and various type III enzymes have the same iron ligands, two histidine and one glutamate, that constitute the 2-His-1carboxylate structural motif (Vaillancourt et al., 2006). 2.3.5.1

The Gentisate Pathway

Gentisate and substituted gentisates serve as the focal point in the aerobic biodegradation of a large number of simple and complex aromatic hydrocarbons such as salicylate,

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3-hydroxybenzoate, 3,5- or 2,5-xylenol or naphthalene. Consequently, the gentisate pathway is distributed throughout the bacterial world. In this pathway, gentisate 1,2-dioxygenase, a member of the cupin superfamily, cleaves the aromatic ring between the carboxyl substituent and the proximal hydroxyl group to yield maleylpyruvate (Crawford et al., 1975) (> Fig. 9). Gentisate 1,2-dioxygenases have been purified and characterized from various Proteobacteria and Actinobacteria (Crawford et al., 1975; Suemori et al., 1993), and even archaea (Fu and Oriel, 1998), and are described as twodomain bicupins (Dunwell et al., 2000). The gentisate 1,2-dioxygenases reported to date have demonstrated reasonably broad substrate tolerance in terms of substitutions on the aromatic ring catalyzing the turnover of a range of alkyl- and halo-substituted gentisates (Harpel and Lipscomb, 1990). Rates similar to that observed with gentisate are reported with C-3 substituted gentisates (methyl, ethyl, 2-propyl, bromo, fluoro), whereas C-4 substituted gentisates are turned over at reduced rates (Crawford et al., 1975; Harpel and Lipscomb, 1990). All three functional groups of gentisate appear to be required for efficient turnover. However derivatives having a substitution of the carboxyl-group by hydroxyl, acetyl, or aldehyde functions are slowly metabolized (Lipscomb and Orville, 1992). Maleylpyruvate produced by gentisate 1,2-dioxygenase is converted to Krebs cycle intermediates via two downstream routes, a direct hydrolytic cleavage to pyruvate and maleate by maleylpyruvate hydrolase (Hopper et al., 1971) or isomerization to fumarylpyruvate and subsequent hydrolytic cleavage to fumarate and pyruvate by fumarylpyruvate hydrolase

. Figure 9 Metabolism of gentisate, homogentisate and salicylate via 1,2-dioxygenolytic cleavage.

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(Crawford and Frick, 1977) (> Fig. 9). In the latter pathway, isomerization of maleylpyruvate to fumarylpyruvate is catalyzed by either a glutathione (GSH)-dependent maleylpyruvate isomerase almost exclusively found in gram negative bacteria (Crawford et al., 1975), or a GSH-independent maleylpyruvate isomerase that has been characterized in various grampositive bacteria (Crawford and Frick, 1977; Shen et al., 2005). Sequence analysis of the GSH-dependent and -independent maleylpyruvate isomerases revealed that the two isomerases were neither homologous nor phylogenetically related (Shen et al., 2005). 2.3.5.2

The Homogentisate Pathway

Homogentisate is the central metabolite formed during degradation of aromatic amino acids phenylalanine and tyrosine in several microorganisms (see 2.2.5). Its further metabolism is initiated by homogentisate 1,2-dioxygenase, which perform the ring cleavage between the acetyl substituent and the proximal hydroxyl group to yield maleylacetoacetate in a manner analogous to that of gentisate 1,2-dioxygenase (Harpel and Lipscomb, 1990; Titus et al., 2000) (> Fig. 9). Like gentisate 1,2-dioxygenase, also homogentisate 1,2-dioxygenase is a type III extradiol dioxygenases of the two-domain bicupins, that usually contain a single active site in one of two domains, with the other domain remaining as a non-catalytic vestigial remnant (Vaillancourt et al., 2004). The downstream catabolism of maleylacetoacetate is analogous to that described for maleylpyruvate in the gentisate pathway (> Fig. 9). It can be hydrolyzed directly to acetoacetate and maleate by maleylacetoacetate hydrolase (Crawford, 1976) or, obviously more commonly, be isomerized by a GSH-independent (Suemori et al., 1996) or GSH-dependent isomerase (Crawford and Frick, 1977) to fumarylacetoacetate which is finally hydrolyzed to acetoacetate and fumarate by fumarylacetoacetate hydrolase (Arias-Barrau et al., 2004; Crawford and Frick, 1977). 2.3.5.3

Direct Cleavage of Salicylate by Salicylate 1,2-Dioxygenase

Recently, a new ring fission dioxygenase which cleaves salicylate between the carboxyl group and the hydroxyl group to form 2-oxohepta-3,5-dienedioate has been described in the naphthalenesulfonate-degrading strain Pseudaminobacter salicylatoxidans BN12 (Hintner et al., 2001). Similarly, 1-hydroxy-2-naphthoate dioxygenase from Nocardioides sp. KP7 (Iwabuchi and Harayama, 1998) involved in the degradation of phenanthrene by this strain was shown to be capable to cleave between a carboxyl and a hydroxyl group, contradicting a generally accepted paradigm that the enzymatic ring fission of the aromatic nucleus by bacteria requires the presence of two hydroxyl groups or one amino and one hydroxyl group (> Fig. 9). In addition to salicylate, salicylate 1,2-dioxygenase also converts gentisate and a wide range of substituted salicylates (Hintner et al., 2001). The deduced amino acid sequence revealed that salicylate-1,2-dioxygenase also belongs to the type III extradiol dioxygenases with a subunit topology characteristic of the bicupin beta-barrel folds (Matera et al., 2008). The crystal structure revealed, however, that this enzyme does not contain the classical 2-His-1carboxylate metal-binding motif but a mononuclear iron center involving three histidine ligands, the iron coordination being completed by water molecules (Matera et al., 2008). The downstream pathway of 2-oxohepta-3,5-dienedioate is still elusive. 2.3.5.4

The 3-Hydroxyanthranilate Pathway

3-Hydroxyanthranilate is a central intermediate of tryptophan degradation via the kynurenine pathway (see 2.2.5) (> Fig. 5) and of the biosynthetic pathway from tryptophan to quinolinate,

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the universal de novo precursor to the pyridine ring of nicotinamide adenine dinucleotide. In this pathway, 3-hydroxyanthranilate 3,4-dioxygenase catalyzes the conversion of 3-hydroxyanthranilate to 2-amino-3-carboxymuconic semialdehyde (Muraki et al., 2003) (> Fig. 7). 3-Hydroxyanthranilate 3,4-dioxygenase also belongs to the type III extradiol dioxygenases, but in contrast to gentisate 1,2-dioxygenase, it is composed of a single cupin domain (Zhang et al., 2005). The ring-cleavage product 2-amino-3-carboxymuconic semialdehyde is decarboxylated to 2-aminomuconic semialdehyde (Muraki et al., 2003), a common intermediate with the 2-aminophenol metabolic pathway (see 2.3.4.4) (> Fig. 7). 2.3.5.5

The 4-Amino-3-Hydroxybenzoate Pathway

A 4-amino-3-hydroxybenzoate 2,3-dioxygenase that catalyzes the ring fission between C2 and C3 yielding 2-amino-5-carboxymuconic semialdehyde has been isolated from Bordetella sp. 10d (Takenaka et al., 2002). This Fe2+ dependent enzyme is highly specific and neither 2-aminophenol, nor its methyl-, hydroxyl- or carboxyl- derivatives, including 3-hydroxyanthranilate are substrates. The deduced amino acid sequence shows significant identity (28%) with 3-hydroxyanthranilate 3,4-dioxygenases indicating that this enzyme also belongs to the cupin superfamily (Murakami et al., 2004). Further metabolism of 2-amino-5-carboxymuconic semialdehyde is assumed to proceed via enzyme catalyzed deamination to 2-hydroxy-5-carboxymuconic semialdehyde followed by spontaneous decarboxylation to yield 2-hydroxymuconic semialdehyde (Orii et al., 2004). Subsequent catabolism of 2-hydroxymuconic semialdehyde occurs via a dehydrogenative route as described above (see 2.3.2) (> Fig. 7).

2.4

CoA Dependent Pathways

The involvement of CoA-dependent reactions in aromatic hydrocarbon degradation is known since decades. However, such an involvement was thought to be restricted to an oxidation of side-chains or reactions funneling ring-cleavage products into the Krebs cycle. The cleavage of the aromatic ring of CoA-substituted derivatives was assumed to be restricted to anaerobic pathways. However, a novel aerobic route for degradation of aromatic hydrocarbons without involvement of dihydroxylated aromatic intermediates was initially reported for phenylacetate degradation in E. coli W (Ferrandez et al., 1998) and P. putida U (Olivera et al., 1998) (> Fig. 10). The initial step of the pathway involves the activation of phenylacetate into phenylacetylCoA by a phenylacetate-CoA ligase (Mohamed, 2000). Like other CoA ligases (see 2.2.4), this enzyme belongs to the AMP-forming acyl-CoA ligases, which catalyze thioesterification via a two-step process in which an acyl-adenosine monophosphate (AMP) intermediate is formed in the first step, followed by formation of the acyl-CoA ester and release of AMP. PhenylacetylCoA is attacked by a ring-oxygenase/reductase (the PaaABCDE gene products), generating a hydroxylated and reduced derivative of phenylacetyl-CoA, probably 1,2-dihydroxy-1,2-dihydrophenylacetyl-CoA, which is not re-oxidized to a dihydroxylated aromatic intermediate, as in other known aromatic pathways (Ismail et al., 2003) (> Fig. 10). Sequence comparisons of the paaABCDE gene products strongly suggest that the oxygenase belongs to the bacterial diiron multicomponent oxygenases family and suggest that PaaACD might constitute the a, b, and y subunits of the heteromultimeric diiron oxygenase component of the oxygenase. PaaB

Aerobic Degradation of Aromatic Hydrocarbons

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. Figure 10 Metabolism of phenylacetate, benzoate and 2-aminobenzoate by CoA-dependent aerobic pathways. Reaction intermediates are shown in brackets. The transformation of 3-hydroxyadipylCoA to 3-oxoadipyl-CoA has been proven to be involved in phenylacetate degradation. Tentative reactions are indicated by a question mark.

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and PaaE may be the effector protein and the oxidoreductase, respectively, that mediate electron transfer from NAD(P)H (Fernandez et al., 2006). Interestingly, although all bacterial diiron multicomponent oxygenases described so far are monooxygenases, the proposed product of the reaction catalyzed by the oxygenase is a dihydrodiol, and therefore this enzyme could be a hydroxylating dioxygenase (Ismail et al., 2003). It has been proposed that 1,2dihydroxy-1,2-dihydrophenylacetyl-CoA, is further metabolized in a complex reaction sequence comprising enoyl-CoA isomerization/hydration, non-oxygenolytic ring opening and dehydrogenation, which is catalyzed by the PaaG and PaaZ gene products. The resulting aliphatic CoA dicarboxylate compound is further catabolized by a b-oxidation-like pathway via b-ketoadipyl-CoA (Ismail et al., 2003) (> Fig. 10) and a b-ketoadipyl-CoA thiolase that catalyses the last step of the phenylacetate catabolic pathway, i.e., the thiolytic cleavage of betaketoadipyl-CoA to succinyl-CoA and acetyl-CoA (Nogales et al., 2007). Also benzoate, a strategic intermediate in aerobic aromatic hydrocarbon metabolism, can be metabolized aerobically via benzoyl-CoA, involving also non-oxygenolytic ring cleavage (Altenschmidt et al., 1993) (> Fig. 10). The benzoate-CoA ligase of B. xenovorans LB400, which also belongs to the AMP-forming acyl-CoA ligases, has been analyzed in detail and shows some activity with 2-aminobenzoate, but is inactive with phenylacetate (Bains and Boulanger, 2007). Benzoyl-CoA is hydroxylated by benzoyl-CoA oxygenase/reductase, a two component benzoyl-CoA dioxygenase, forming 2,3-dihydro-2,3-dihydroxybenzoyl-CoA (Zaar et al., 2004) (> Fig. 10). Benzoyl-CoA dioxygenase is composed of an iron-sulfur-flavoprotein reductase (BoxA) and an oxygenase (BoxB) which shows low similarity to PaaA, the supposed a subunit of the heteromultimeric diiron phenylacetyl-CoA oxygenase. The dihydrodiol is the substrate for ring fission catalyzed by dihydrodiol lyase (BoxC) (Gescher et al., 2005), a member of the enoyl-CoA hydratase/isomerase superfamily. This homodimeric enzyme does not require oxygen and catalyzes the transformation to 3,4-dehydroadipyl-CoA semialdehyde. The latter intermediate is subsequently oxidized by 3,4-dehydroadipyl-CoA semialdehyde dehydrogenase (BoxD) to 3,4-dehydroadipyl-CoA (Gescher et al., 2006) (> Fig. 10). The further metabolism is thought to lead to 3-oxoadipyl-CoA, which is finally cleaved into succinyl-CoA and acetyl-CoA (Zaar et al., 2004). An on the first view similar pathway has also been reported for the aerobic metabolism of anthranilate (2-aminobenzoate) via 2-aminobenzoyl-CoA (Altenschmidt and Fuchs, 1992). Even though thioesterification is catalyzed by a 2-aminobenzoate CoA ligase with similarity to benzoate CoA ligase, oxygenation is catalyzed by a 2-aminobenzoyl-CoA monooxygenase/ reductase rather than a diiron oxygenase (Buder and Fuchs, 1989). This enzyme catalyzes both monooxygenation and hydrogenation of 2-aminobenzoyl-CoA to form 2-amino-5-oxocyclohex-1-enecarboxyl-CoA via 2-amino-5-oxocyclohex-1,3-dienecarboxyl-CoA (Schuhle et al., 2001) (> Fig. 10). Sequence analysis revealed that the N-terminal part shows similarities to single component flavin monooxygenases and the C-terminal part to NADH dependent, flavin-containing oxidoreductases of the old-yellow-enzyme type (Schuhle et al., 2001). Further metabolism is assumed to proceed by b-oxidation, however, the metabolic pathway remains to be elucidated. Interestingly, novel CoA dependent pathways are still being discovered. In Streptomyces WA-46 salicylate was described to be subject to initial thioesterification and the formed salicylyl-CoA hydroxylated by salicylyl-CoA 5-hydroxylase to gentisyl-CoA (Ishiyama et al., 2004). Gentisyl-CoA was supposed to spontaneously decompose to CoA and gentisate, which was subject to ring-cleavage by a gentisate 1,2-dioxygenase. However, also gentisyl-CoA thioesterases were recently described (Zhuang et al., 2004).

Aerobic Degradation of Aromatic Hydrocarbons

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4

Research Needs

Various main routes for microbial aerobic degradation of aromatic hydrocarbons are known. However, novel catabolic pathways are still being discovered, indicating the broad and still poorly understood diversity of microbial capabilities. Also the links between aerobic and anaerobic degradation of aromatic hydrocarbons are poorly described. Changing environments are common for microorganisms, especially bacteria, therefore oxygen availability may control the way a particular aromatic hydrocarbon is being degraded. In this context, CoA dependent pathways may play an important role. Another important research need is to broaden the knowledge on the range and type of peripheral reactions that microorganisms can perform. Interestingly, even the metabolism of abundant aromatics, such as the amino acid tryptophan is still poorly understood. In view of the impressive variety of natural aromatic products, especially those produced by plants, this unexplored diversity can be an important source of enzymes for transformation to valuable products. Special efforts should be directed towards a better understanding of O-demethylation reactions, because the number of methoxylated aromatic hydrocarbons known is far greater than the methoxylated aromatic hydrocarbons known to be degraded. It would be also important to better understand why some compounds are degraded by different peripheral or central pathways. For example, benzoate can be degraded by the classical ortho ring cleavage pathway, but also by a CoA dependent pathway; toluene can be degraded by direct oxygenation of the aromatic ring and also by oxidation of the methyl substituent; catechol and protocatechuate can be degraded through ortho or meta ring cleavage pathways. Is the particular pathway controlled by physiological constraints at the cell level? Is it controlled at the species or population level or by environmental factors, such as oxygen or iron availability, as suggested for benzoate degradation pathways? As it is thoroughly demonstrated in this chapter, an impressive diversity of oxygenases plays a significant role at different stages in aerobic aromatic hydrocarbon degradation. Several aspects concerning oxygenases should be addressed. Substrate specificity is a key to allow these enzymes to use different compounds as substrates. Narrow specificity decreases the impact of a particular oxygenase in aerobic degradation whereas broad specificity, in principle, provides an advantage allowing the microorganism to degrade a wider range of (potential) carbon and energy sources. However, this potential advantage contrast with problems associated to deadend product formation and, more important, intermediate misrouting. Additional biochemical studies, especially those related with regioselectivity, and genetic studies, i.e., inducer of the genes encoding oxygenases, are clearly required. In addition, ongoing microbial genome sequencing projects clearly indicate the presence of sequences putatively encoding oxygenases that do not match, or cannot be associated with the pathways, which have been already reported. Although a number of these putative sequences may be related to biosynthesis or even degradation of non-aromatic compounds, it is highly expected that a significant fraction of them, would be involved in aerobic aromatic hydrocarbon degradation. Metabolic reconstruction studies linking in vivo with in silico catabolic properties and transcriptional studies would help to address this point. Moreover, as stated in > Chapter 5, Vol. 2, Part 2, most of the current knowledge on biochemistry and genetics of aromatic aerobic microbial metabolism has been obtained with bacterial strains isolated by traditional culture dependent approaches. Taking into account the significant increase in knowledge on strategies to degrade aromatics, which is still obtained by

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new isolates, it seems obvious that our current knowledge cover only a small proportion of the broad microbial degradative potential.

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5 Aerobic Degradation of Chloroaromatics D. H. Pieper1 . B. Gonza´lez 2 . B. Ca´mara1 . D. Pe´rez-Pantoja2 . W. Reineke3 1 Biodegradation Research Group, Division of Microbial Pathogenesis, HZI – Helmholtz Research Centre for Biotechnology, Braunschweig, Germany [email protected] 2 Departamento de Gene´tica Molecular y Microbiologı´a, Facultad de Ciencias Biolo´gicas, NM-EMBA, CASEB, P. Universidad Cato´lica de Chile, Santiago, Chile 3 Chemical Microbiology, Bergische Universita¨t Wuppertal, Wuppertal, Germany 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 840

2 2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.1.5 2.1.6 2.1.7

2.2.4 2.2.5 2.2.6 2.3

Metabolism of Chloroaromatics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 840 Peripheral Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 840 Rieske Non-Heme Iron Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 842 Oxygenolytic Dehalogenations by Rieske Non-Heme Iron Oxygenases . . . . . . . . . . . 842 Soluble Diiron Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 843 Single Component Flavoprotein Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 844 Initial Side-Chain Processing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 845 Hydrolytic Dehalogenation of 4-Chlorobenzoate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 847 Reactions Leading to (Chloro)Benzoquinols and (Chloro) Hydroxybenzoquinols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 848 Degradation of Chlorocatechols and Chloroprotocatechuates . . . . . . . . . . . . . . . . . . . . 850 Metabolism of Chlorocatechols via the 3-Oxoadipate Pathway . . . . . . . . . . . . . . . . . . . 850 The Chlorocatechol Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 853 The Alternative Pathway of 3-Chlorocatechol Degradation in Rhodococcus opacus 1CP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 855 The Alternative Pathway of 4-Halocatechol Degradation . . . . . . . . . . . . . . . . . . . . . . . . . 855 Chlorocatechol Degradation via the Meta-Cleavage Pathway . . . . . . . . . . . . . . . . . . . . . . 856 Degradation of 5-Chloroprotocatechuate via a Meta-Cleavage Pathway . . . . . . . . . . 857 Ring Cleavage of (Chloro)Benzoquinols and (Chloro)Hydroxybenzoquinols . . . . 858

3

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 859

2.2 2.2.1 2.2.2 2.2.3

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_61, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Microorganisms are key players in the global carbon cycle. In addition, it appears that most xenobiotic industrial chemicals can be degraded by microorganisms, either by a combination of cometabolic steps or by serving as growth substrate, leading to the mineralization of at least part of the molecule. Here, we present the principles of the microbial aerobic degradation of chloroaromatic compounds. The so-called peripheral sequences of the oxidative degradation of chloroaromatic compounds in aerobic bacteria yield central intermediates with a diphenolic structure such as catechols or hydroxybenzoquinols. These compounds are subsequently cleaved by enzymes that use molecular oxygen and further metabolized by central pathway sequences. The broad variety of mechanisms resulting in dechlorination that occur in these peripheral or central sequences is specifically discussed.

1

Introduction

Chlorinated hydrocarbons comprise a large spectrum of compounds that are or have been of enormous industrial and economic importance. The introduction of chlorine atom(s) into a hydrocarbon significantly influences its physicochemical and biochemical properties and the tendency for bioaccumulation and environmental persistence. Acting in combination with possible (eco)toxicological effects, these properties have pushed the chlorochemistry into the focus of considerable debate and governmental regulatory action. For decades, it is known that microorganisms have the capability to mineralize various chlorinated hydrocarbons. Under anaerobic conditions, chlorinated hydrocarbons can function as alternative electron acceptors in a process termed dehalorespiration. Under aerobic conditions, chlorinated hydrocarbons can function as carbon and energy source, which necessitates dechlorination. This chapter deals with the aerobic degradation and the use of the chloroaromatics as carbon and energy source involving elimination of chloride from the substrate or metabolites.

2

Metabolism of Chloroaromatics

Aerobic microorganisms usually initiate degradation by activation of the aromatic nucleus through oxygenation reactions. A few central intermediates such as catechols, protocatechuate, gentisate, and hydroxybenzoquinols are produced by the introduction of hydroxyl groups, usually in ortho- or para-position to one another (peripheral reactions). These intermediates are subject to oxygenolytic ring cleavage followed by the channeling of the ring-cleavage products into the central metabolism (See > Chapter 4, Vol. 2, Part 2). Despite the fact that various specific dehalogenating enzymes have been identified, microorganisms capable of mineralizing chloroaromatics often employ peripheral reactions, which have their function in the degradation of naturally occurring aromatics such as benzoate, salicylate, or biphenyl (See > Chapter 26, Vol. 2, Part 5) (Reineke, 2001). These enzyme systems are predominantly of relaxed-substrate specificity and tolerate lower chlorinated substrate analogs.

2.1

Peripheral Reactions

A broad set of peripheral pathways for chloroaromatic degradation has been described, mainly responsible for transforming chloroaromatics to central catechol or benzoquinol intermediates. > Figure 1 gives an overview on reactions leading to (chloro) catechols.

. Figure 1 Overview of peripheral reactions in the degradation of chloroaromatics channeling to catechols. (1) Rieske non-heme iron oxygenases (1a) and dehydrogenases (1b) involved in (chloro)benzene and (chloro)benzoate metabolism; (2) aromatic monooxygenases (soluble diiron monooxygenases); (3) phenol hydroxylases (soluble diiron monooxygenases or single component flavin monooxygenases); (4) salicylate 1-hydroxylases (single component flavin monooxygenases); (5) a-ketoglutarate-dependent dioxygenases; (6) 2,4,5-trichloro- and 2,4-dichlorophenoxyacetate monooxygenases; (7) oxidation of side chains (xylene monooxygenases, 7a; benzylalcohol dehydrogenases, 7b; benzaldehyde dehydrogenases, 7c).

Aerobic Degradation of Chloroaromatics

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5 2.1.1

Aerobic Degradation of Chloroaromatics

Rieske Non-Heme Iron Oxygenases

The so called Rieske non-heme iron oxygenases are one of the key families of enzymes important for aerobic activation and thus degradation of aromatics such as benzoate, benzene, toluene, phthalate, naphthalene, or biphenyl (Gibson and Parales, 2000) (see > Fig. 1, reaction 1). These multicomponent enzyme complexes, composed of a terminal oxygenase component and different electron transport proteins, usually catalyze the incorporation of two oxygen atoms into the aromatic ring to form arene-cis-dihydrodiols, a reaction that is followed by a dehydrogenation catalyzed by cis-dihydrodiol dehydrogenases to give (substituted) catechols. Comparison of the amino acid sequences of the terminal oxygenase a-subunits revealed that they form a family of diverse but evolutionarily related sequences. Although none of the enzymes is completely specific, a broad correlation between the grouping in toluene/biphenyl, naphthalene, benzoate, or phthalate families, and the native substrates oxidized by the family members can be observed (Gibson and Parales, 2000). Enzyme engineering studies of biphenyl, benzene, chlorobenzene, and naphthalene dioxygenases showed that the a-subunit of the terminal oxygenase determines substrate specificity and that only slight differences in the amino acid sequence can be associated with dramatic changes in substrate specificity or regioselectivity (Beil et al., 1998; Furukawa et al., 2004). However, as a general rule Rieske non-heme iron oxygenases can transform lower chlorinated substrate analogs and dioxygenation is usually directed to carbon atoms proximal to substituents.

2.1.2

Oxygenolytic Dehalogenations by Rieske Non-Heme Iron Oxygenases

Different bacteria capable of degrading 2-chlorobenzoate have been described. All these organisms catalyze a 1,2-dioxygenation such that one of the vic-hydroxyl groups in the cisdihydrodiol is bound to the same carbon as the chloro-substituent. From such an unstable vic-dihydrodiol, the chloro-substituent is spontaneously eliminated to form catechols (> Fig. 2). Two distinct 2-chlorobenzoate-degrading dioxygenase systems have been described. The two-component 2-halobenzoate 1,2-dioxygenases (oxygenase consisting of a- and b-subunits and a reductase as the one from strain Burkholderia cepacia 2CBS (Fetzner et al., 1992)), are similar to two-component toluate and benzoate 1,2-dioxygenases of the Rieske non-heme iron oxygenases. They are characterized by their high activity against 2-halosubstituted benzoates but have negligible activity with 4-chloro-, or 2,5-dichlorobenzoate. In contrast, the broad

. Figure 2 Oxygenolytic dehalogenation of 2-chlorobenzoate by Rieske non-heme iron oxygenases.

Aerobic Degradation of Chloroaromatics

5

specificity 2-chlorobenzoate dioxygenase of P. aeruginosa strain 142 is a three-component dioxygenase system (oxygenase consisting of a- and b-subunits, ferredoxin and reductase) (Romanov and Hausinger, 1994), the a-subunit of which exhibits only 22% sequence identity with that of strain 2CBS, but a significant similarity (42%) to salicylate 5-hydroxylase NagG from Pseudomonas sp. strain U2. Thus, 2-chlorobenzoate dioxygenases are functionally similar, but represent two different lineages with distinct activities. Oxidative dehalogenation is not restricted to 2-halobenzoate 1,2-dioxygenases but has also been described for tetrachlorobenzene dioxygenase TecA of Ralstonia sp. PS12 (Beil et al., 1998), biphenyl 2,3-dioxygenase BphA of Burkholderia xenovorans LB400 (Seeger et al., 1995), or 3-chlorobenzoate 4,5-dioxygenase CbaA from Comamonas testosteroni BR60 (Nakatsu and Wyndham, 1993). Dehalogenation always involves dioxygenolytic attack on a chlorosubstituted carbon atom and its unsubstituted neighbor to give unstable intermediates, which spontaneously rearrange with elimination of chloride. However, in all cases mentioned here, only higher chlorinated substrate analogs are dechlorinated. CbaA catalyzes the 4,5-dioxygenation of 3,4-dichlorobenzoate resulting in an unstable dihydrodiol, which spontaneously eliminates chloride to form 5-chloroprotocatechuate, whereas 4,5-dioxygenation of 3-chlorobenzoate yields 5-chloroprotocatechuate after dehydrogenation. TecA catalyzes the dehalogenation of 1,2,4,5-tetrachlorobenzene to form 3,4,6-trichlorocatechol, whereas lower chlorinated benzenes were transformed to the corresponding dihydrodiols. BphA catalyzes the dehalogenation of 2,20 - or 2,40 -dichlorobiphenyl among others (See > Chapter 26, Vol. 2, Part 5). For both TecA and BphA, amino acid residues crucial for dehalogenation were identified (Beil et al., 1998; Furukawa et al., 2004).

2.1.3

Soluble Diiron Monooxygenases

Enzymes attacking the nonactivated benzene nucleus by monooxygenation belong to an evolutionary-related family of soluble diiron monooxygenases (Leahy et al., 2003) that are enzyme complexes consisting of an electron transport system comprising a reductase (and in some cases a ferredoxin), a catalytic effector protein, and a terminal hydroxylase with a (abg)2 quaternary structure and a diiron center contained in each a-subunit. The monooxygenases are classified according to their a-subunits, which are assumed to be the site of substrate hydroxylation, into four different phylogenetic groups, comprising the (multicomponent) phenol hydroxylases, and the four-component alkene/aromatic monooxygenases (Leahy et al., 2003). The multicomponent phenol hydroxylases (> Fig. 1, reaction 3) such as the phenol hydroxylase of Pseudomonas sp. CF600 share the capability to hydroxylate phenol and methylsubstituted derivatives (Shingler et al., 1992), and a few enzymes of this group, such as phenol hydroxylase of P. stutzeri OX1, also have been shown to hydroxylate the nonactivated benzene nucleus and thus can catalyze two sequential hydroxylations (Cafaro et al., 2004). The available information on the transformation of chlorophenols by multicomponent phenol hydroxylases is insufficient; however, preliminary data indicate that such a capability is spread among this group of enzymes (Teramoto et al., 1999). The four-component alkene/aromatic monooxygenases comprise enzymes that oxidize nonhydroxylated compounds (Leahy et al., 2003) (> Fig. 1, reaction 2). Toluene 4-monooxygenase of P. mendocina KR1 also transforms chlorobenzene with 4-chlorophenol as product (Fishman et al., 2005). Regioselectivity of attack is mainly controlled by the a-subunit of the enzyme, but

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Aerobic Degradation of Chloroaromatics

also by the effector protein (Mitchell et al., 2002), in such a way that the enzyme could be modulated also to produce 2-chloro- and 3-chlorophenol. Tbc2 toluene 2-monooxygenase of strain Burkholderia cepacia JS150 was shown to transform chlorobenzene into 2-chlorophenol (Kahng et al., 2001).

2.1.4

Single Component Flavoprotein Monooxygenases

The oxidation of various phenolic aromatic compounds is catalyzed by single-component flavoprotein monooxygenases (class A flavin monooxygenases (van Berkel et al., 2006)). These enzymes catalyze hydroxylation in ortho- or para-position to the preexisting hydroxyl-group (See > Chapter 4, Vol. 2, Part 2) and usually display a narrow substrate specificity (van Berkel et al., 2006). Only a few of these enzymes have been analyzed for their capability to transform chlorinated substrate analogues. Single-component salicylate 1-hydroxylases (> Fig. 3) have been reported to catalyze the transformation of salicylate to catechol (> Fig. 1, reaction 4), and enzymes analyzed in this aspect were capable of transforming chlorinated salicylates into the respective chlorocatechols with high activity (Bosch et al., 1999; Ca´mara et al., 2007). In accordance with these properties, mineralization of chlorosalicylates by natural or engineered organisms involving salicylate 1-hydroxylase has been reported (Ca´mara et al., 2007; Lehrbach et al., 1984). Another group of single component flavin monooxygenases, termed phenol hydroxylases (> Fig. 1, reaction 3), has been described from phenol degrading Pseudomonas strains, among them PheA from Pseudomonas sp. strain EST1001, which transforms phenol and 3-methylphenol (Nurk et al., 1991). This substrate specificity significantly differs from that described for a group of enzymes termed 2,4-dichlorophenol hydroxylases and involved in the degradation of 2,4-D or 2,4-dichlorophenol via 3,5-dichlorocatechol, with which phenol hydroxylases share 50% sequence identity (> Fig. 3). In contrast to the substrate specificity of phenol hydroxylases, all the five characterized dichlorophenol hydroxylases, TfdB of Burkholderia cepacia 2a (Beadle and Smith, 1982), ClpB from Defluvibacter lusatiensis S1 (Makdessi and Lechner, 1997), TfdBa from Bradyrhizobium sp. RD5-C2 (Huong et al., 2007), and both TfdBI and TfdBII dichlorophenol hydroxylases from C. necator JMP134 (Ledger et al., 2006), are highly active with 2,4-dichloro-, 4-chloro-2methylphenol and 4-chlorophenol, but activity with phenol and 3-chlorophenol was either low or absent. Despite the obvious similarity in substrate specificity, these dichlorophenol hydroxylases are just distantly related. Various 2,4-D degraders contain dichlorophenol hydroxylases sequences highly related to that of B. cepacia 2a or TfdBII from strain JMP134 (e.g., A. denitrificans EST4002, D. acidovorans P4a, Burkholderia sp. RASC or V. paradoxus TV1). In contrast, the closest homolog of TfdBI from strain JMP134 is the enzyme of D. lusatiensis (only 76% identity). However, partial amino acid sequences indicate that enzymes highly similar to TfdBI are present in other 2,4-D degrading b-proteobacterial strains (Vallaeys et al., 1999). In addition to ClpB from D. lusatiensis and TfdBa from Bradyrhizobium sp., a third lineage of dichlorophenol hydroxylases represented by TfdB of S. herbicidovorans MH (Mu¨ller et al., 2004) has been observed in a-proteobacteria. All these three lineages have only approximately 60% amino acid identity (> Fig. 3) and similar identity to TfdB proteins from b-proteobacteria, indicating an ancient divergence and evolution without recent horizontal gene transfer in a-proteobacteria.

Aerobic Degradation of Chloroaromatics

5

. Figure 3 Dendrogram showing the relatedness of single component flavin monooxygenases (2,4-dichlorophenol hydroxylase, TfdB, ClpB; phenol hydroxylase, PheA; salicylate 1-hydroxylase, SalA, NahG, NahW; pentachlorophenol monooxygenase, PcpB; 2-hydroxybiphenyl 3-monooxygenase, HbpA; chlorobenzoquinol monooxygenase, ChqA; unknown function, CphC-II).

2.1.5

Initial Side-Chain Processing

2.1.5.1

Oxidation of Methylgroups

Aromatic compounds that bear alkyl substituents on the aromatic ring may undergo oxidation of the side chain before ring activation. The most intensely described example is the TOL plasmid encoded degradation pathway of toluene and xylenes by Pseudomonas putida mt-2, which proceeds via benzoate or the respective methylbenzoates, and catechol or methylcatechols as intermediates (Worsey and Williams, 1975) (> Fig. 1, reactions 7). The respective enzymes are encoded by the xyl genes, which are organized in two functional units localized on

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Aerobic Degradation of Chloroaromatics

the TOL plasmid pWW0 (Greated et al., 2002). The upper pathway encodes three enzymes, xylene monooxygenase, benzylalcohol dehydrogenase, and benzaldehyde dehydrogenase, that oxidize toluene and xylenes to benzoate and toluates, respectively. The degree of transformation of chlorotoluenes by xylene monooxygenase depends on the position of the chlorine substituent. The substrate analogs 3-chloro- and 4-chlorotoluene are transformed at high rates, while no or only low activity has been found with other chlorotoluenes. Substituents in ortho position impaired substrate binding (Brinkmann and Reineke, 1992). Benzylalcohol and benzaldehyde dehydrogenases show broader specificities. But again, a chlorine substituent in ortho position leads to a drastic decrease in the substrate conversion. Transfer of the TOL plasmid into strains capable of mineralizing chlorocatechols (see below) allowed the isolation of 3-chloro- and 4-chlorotoluene-degrading organisms (Brinkmann and Reineke, 1992). 2.1.5.2

a-Ketoglutarate-Dependent Dioxygenases

Bacterial metabolism of 2,4-D and MCPA is initiated by the cleavage of the side chain, resulting in the formation of 2,4-dichloro- and 4-chloro-2-methylphenol, respectively (> Fig. 1, reaction 5). In C. necator JMP134 (pJP4), the degradation of 2,4-D is catalyzed by 2,4-dichlorophenoxyacetate/a-ketoglutarate dioxygenase, TfdA (Fukumori and Hausinger, 1993). TfdA belongs to the large superfamily of a-ketoacid-dependent, mononuclear nonheme iron oxygen activating enzymes that catalyze the oxidation of aliphatic C–H bonds (Schofield and Zhang, 1999). During 2,4-D transformation, one oxygen atom is involved in the oxidative decarboxylation of a-ketoglutarate, whereas the other oxygen atom is introduced into the b-carbon of the aliphatic side chain to produce an unstable hemiacetal intermediate that decomposes to glyoxylate, and 2,4-DCP (Fukumori and Hausinger, 1993). TfdA is able to hydroxylate several phenoxyacetates, including 2,4,5-trichlorophenoxyacetate (2,4,5-T) and MCPA; however, 2,4-D is the preferred substrate. Representatives of tfdA genes have been described thus far from a, b, and g subgroups of the proteobacteria (McGowan et al., 1998). Sequence alignment among b-proteobacterial representatives reveals the presence of three distinct classes of tfdA gene sequences with slight variations in each class (> Fig. 4); however, all share >80% sequence identity and all are capable of transforming several phenoxyacetate compounds. tfdA-like genes were also observed in 2,4-D degrading a-proteobacteria belonging to the Bradyrhizobium-AgromonasNitrobacter-Afipia cluster (Itoh et al., 2002), and these tfdAa sequences show 56–60% identity to the b-proteobacterial counterparts. tfdA-like genes were also detected in 2,4-D degrading sphingomonads showing 46–51% of nucleotide sequence identity with either tfdA or tfdAagenes (Itoh et al., 2004) (> Fig. 4). Dichlorprop ((RS)-2-(2,4-dichlorophenoxy)propionate) and mecoprop ((RS)-2(4-chloro-2-methylphenoxy)propionate) are chiral molecules that are poor substrates for the TfdA enzymes already mentioned. Experiments with pure enantiomers of mecoprop revealed that S. herbicidovorans MH and D. acidovorans MC1 can degrade both enantiomers. In both organisms, enantioselective a-ketoglutarate-dependent dioxygenases termed SdpA and RdpA were observed (Mu¨ller et al., 2006; Westendorf et al., 2003) with the SdpA proteins (64% sequence identity) being highly specific in transforming only the (S)-enantiomers of mecoprop and dichlorprop and the (identical) RdpA proteins active only against the (R)-enantiomers. Like TfdA, SdpA and RdpA are a-ketoglutarate-dependent dioxygenases; however SdpA and RdpA share only 25% of protein sequence identity and only less than 35% of identity with previously characterized TfdA proteins (> Fig. 4).

Aerobic Degradation of Chloroaromatics

5

. Figure 4 Dendrogram showing the relatedness of 2,4-D/a-ketoglutarate dioxygenases (TfdA), (S)-mecoprop/and (R)-mecoprop/a-ketoglutarate dioxygenases (SdpA and RdpA, respectively). TauD taurine a-ketoglutarate dioxygenase from E. coli is shown as outgroup.

2.1.6

Hydrolytic Dehalogenation of 4-Chlorobenzoate

Various Arthrobacter, Pseudomonas and Alcaligenes strains degrading 4-chlorobenzoate (4CB) by a pathway involving an initial dehalogenation event have been described. The 4-chlorobenzoate dehalogenase system consists of three distinct enzymes, a 4CBcoenzyme A (CoA) ligase, 4CB-CoA dehalogenase, and 4-hydroxybenzoate (4HB)-CoA thioesterase (> Fig. 5) encoded by the fcbA, fcbB and fcbC genes, respectively. In 4CB degrading actinobacteria, a fcbABC gene order is conserved (Schmitz et al., 1992), whereas 4CB degrading proteobacterial strains exhibit a fcbBAC gene order (Savard et al., 1992). In some cases, the gene cluster also encodes putative transport proteins between the fcbB and fcbA genes. In addition to differences in the gene order, significant differences in protein sequence were observed. Actinobacterial 4CB-CoA ligases are highly similar (>95% sequence identity), but differ significantly (38–44% identity) from proteobacterial 4CB-CoA ligases; however, all ligases share significant sequence similarity with proteins catalyzing similar chemistry in the boxidation pathway (Babbitt et al., 1992). The following dehalogenation by 4CB-CoA dehalogenase forms 4-hydroxybenzoyl-CoA in a hydrolytic substitution reaction (Liang et al., 1993).

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. Figure 5 Degradation of 4-chlorobenzoate involving hydrolytic dehalogenation.

This group of enzymes utilizes a unique form of catalysis in which an active site carboxylate bonds to C-4 of the benzoyl ring of the bound substrate to form a Meisenheimer-like complex. Expulsion of the chloride from the Meisenheimer complex with concomitant rearomatization of the benzoyl ring generates an arylated enzyme as the second reaction intermediate (Dong et al., 2002). Hydrolysis of the arylated enzyme occurs by addition of a water molecule to the acyl carbonyl carbon to form a tetrahedral intermediate, which expels the hydroxylbenzoyl group to generate the catalytic carboxylate residue and form 4-hydroxybenzoyl-CoA. As observed for the ligases, the actinobacterial dehalogenases are highly similar (>98% sequence identity) but differ significantly (approximately 50% sequence identity) from proteobacterial dehalogenases. The last reaction step to form 4HB is carried out by the 4HB-CoA thioesterase. The absence of serine, cysteine, or histidine catalytic residues in the Pseudomonas sp. strain CBS-3 enzyme had distinguished this protein from previously characterized thioesterases. However, crystallographic investigation (Benning et al., 1998) revealed a high similarity of its threedimensional structure to that of b-hydroxydecanoyl thiol ester dehydrase from E. coli. Both enzymes contained the so-called Hotdog fold, which now was found to be shared by numerous thioesterases and dehydrase proteins (Dillon and Bateman, 2004). An active site aspartate was suggested to participate in nucleophilic catalysis (Zhuang et al., 2002). Other proteobacterial 4HB-CoA thioesterases belong to the same subfamily of Hotdog fold proteins, whereas the respective thioesterases from Arthrobacter strains do not share significant sequence similarity with the proteobacterial counterparts. However, they also belong to the same superfamily and an active site glutamate was suggested as catalytic nucleophile (Dillon and Bateman, 2004).

2.1.7

Reactions Leading to (Chloro)Benzoquinols and (Chloro) Hydroxybenzoquinols

The degradation of various, specially higher chlorinated aromatic pollutants, proceeds via (chloro)benzoquinols or (chloro)hydroxybenzoquinols as central intermediates (> Fig. 6). 2.1.7.1

Degradation of Lindane

One of the most important xenobiotic compounds metabolized through (chloro)benzoquinol is lindane or g-hexachlorocyclohexane (g-HCH), the biodegradation of which has been mainly studied in Sphingobium japonicum UT26 (Endo et al., 2005) (> Fig. 6). However, other g-HCH degrading strains have recently been found to possess nearly identical genes for degradation (Nagata et al., 2007). LinA g-HCH dehydrochlorinase catalyzes two steps of dehydrochlorination to 1,3,4,6-tetrachloro-1,4-cyclohexadiene (1,4-TCDN) via

Aerobic Degradation of Chloroaromatics

5

. Figure 6 Catabolic pathways leading to (chloro)benzoquinols and (chloro)hydroxybenzoquinols as central intermediates. The metabolism of g-hexachlorocyclohexane by S. japonicum UT26, of pentachlorophenol by S. chlorophenolicum ATCC39723, of 2,4,6-trichlorophenol by C. necator JMP134 and of 2,4,5-trichlorophenol by B. phenoliruptrix AC1100 are shown. Unstable intermediates are enclosed in brackets. Broken arrows indicate reactions of the lower pathway detailed in > Fig. 10.

g-pentachlorocyclohexene (Nagata et al., 2007). 1,4-TCDN is further transformed by LinB 1,4-TCDN chlorohydrolase: a reaction competing with spontaneous rearrangement to the dead-end 1,2,4-trichlorobenzene product. Unlike LinA, which seems to be a unique dehydrochlorinase, LinB belongs to the haloalkane dehalogenase family and catalyzes two successive hydrolytic dehalogenations to form 2,5-dichloro-2,5-cyclohexadiene-1,4-diol via 2,4,5-trichloro2,5-cyclohexadiene-1-ol (Nagata et al., 2007). Further metabolism of 2,5-dichloro-2,5cyclohexadiene-1,4-diol is achieved by LinC or LinX 2,5-dichloro-2,5-cyclohexadiene-1,4-diol dehydrogenase, members of the short-chain alcohol dehydrogenase superfamily (Nagata et al., 1994), forming 2,5-dichloro-p-benzoquinol, which is subject to a reductive dechlorination by 2,5-dichloro-p-benzoquinol dechlorinase (LinD). LinD belongs to the glutathione transferases family, and catalyzes a rapid dechlorination to chlorobenzoquinol and a subsequent slow conversion to further dechlorinate and forms benzoquinol (Miyauchi et al., 1998). 2.1.7.2

Transformation of Chlorinated Phenols and Phenoxyacetates to (Chloro)Benzoquinols and (Chloro)Hydroxybenzoquinols

Like HCH, pentachlorophenol (PCP) is degraded through (chloro)benzoquinol and the pathway has been elucidated in detail in Sphingobium chlorophenolicum ATCC 39723 (Crawford et al., 2007) (> Fig. 6). The oxidation of PCP is catalyzed by PCP 4-monooxygenase

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(PcpB), a flavin monooxygenase with functional domains similar to those of other bacterial flavoprotein monooxygenases specific to phenolic substrates (see > Fig. 3). PcpB can catalyze attack on a broad range of substituted phenols, hydroxylating the para position and removing halogen, nitro, amino, and cyano groups (Xun et al., 1992a). The generated tetrachloro-pbenzoquinone is reduced to tetrachloro-p-benzoquinol by a NADPH-dependent tetrachlorobenzoquinone reductase (PcpD) (Dai et al., 2003). The following TeCH reductive dehalogenase (PcpC)-catalyzed reactions, reductive dehalogenation to 2,3,6-trichloro-p-benzoquinol and then to 2,6-dichloro-p-benzoquinol (DiCH) (Xun et al., 1992b) resemble those catalyzed by LinD of the HCH degradation pathway. However, in contrast, two chlorine atoms remain on the aromatic ring before ring-cleavage. The broadly used herbicide 2,4,5-T is also metabolized through the (chloro)hydroxybenzoquinol pathway, which has been well studied in Burkholderia phenoliruptrix AC1100 (Hu¨bner et al., 1998). The transformation to 2,4,5-trichlorophenol (2,4,5-TCP) is performed by a twocomponent monooxygenase (TftAB) (Danganan et al., 1995), which exhibits strong homology to terminal oxygenase subunits of benzoate and toluate dioxygenases and thus contrasts the degradation of 2,4-D, which is typically catalyzed by 2,4-D/a-ketoglutarate dioxygenases (see Section 2.1.5.2). However, 2,4-D monooxygenase genes (cadAB) with significant identity (approximately 45% on the amino acid level) with TftAB of B. phenoliruptrix AC1100 were observed in Bradyrhizobium sp. strain HW13 capable of degrading 2,4-D (Kitagawa et al., 2002). The substrate profiles of the 2,4,5-Toxygenase and 2,4-D oxygenase are similar (Danganan et al., 1995; Kitagawa et al., 2002) with high activity against 2,4-D and 2,4,5-T. Additional experiments indicated the presence of respective genes in various 2,4-D degrading Sphingomonas (B6–10, TFD26, TFD44) and Bradyrhizobium (HWK12, BTH) strains (Kitagawa et al., 2002). Further metabolism of 2,4,5-TCP (> Fig. 6) is achieved by TftCD 2,4,5-TCP monooxygenase, a class D monooxygenase (van Berkel et al., 2006) comprising a FADH2-utilizing monooxygenase (TfdD) and a NADH:FAD oxidoreductase (TfdC). TfdC supplies FADH2 to TftD (Gisi and Xun, 2003) converting 2,4,5-TCP via 2,5-dichlorobenzoquinol to 5-chloro-2hydroxybenzoquinol. 5-Chloro-2-hydroxybenzoquinol is dechlorinated to hydroxybenzoquinone by a dechlorinase (TftG) and then reduced to hydroxybenzoquinol by a quinone reductase the gene of which is presently unknown. A similar pathway has been reported for the degradation of 2,4,6-trichlorophenol (2,4,6TCP) in C. necator JMP134 (> Fig. 6). In this bacterium, 2,4,6-TCP monooxygenase (TcpAwith 65% sequence identity to TftD) catalyzes the oxidative conversion of 2,4,6-TCP to 2,6-dichlorobenzoquinone, followed by hydrolytic dechlorination to produce 6-chloro-2-hydroxybenzoquinone (Xun and Webster, 2004). Like TftC, TcpX NADH:FAD oxidoreductase provides FADH2 for TcpA catalysis (Matus et al., 2003). 6-Chloro-2-hydroxybenzoquinone is reduced to 6chlorohydroxybenzoquinol by a quinone reductase (TcpB) (Belchik and Xun, 2008). Recently, evidence was accumulated that also 2,4-dichlorophenol (Ferraroni et al., 2005) and 4-chlorophenol (Nordin et al., 2005) can be degraded via (chloro)hydroxybenzoquinols as intermediate. The downstream catabolism of (chloro)hydroxybenzoquinols is described in Section 2.3.

2.2

Degradation of Chlorocatechols and Chloroprotocatechuates

2.2.1

Metabolism of Chlorocatechols via the 3-Oxoadipate Pathway

The chromosomally encoded 3-oxoadipate pathway is widely distributed in soil bacteria, specifically proteobacteria and actinobacteria (See > Chapter 39, Vol. 2, Part 6). The function

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of the catechol branch of this pathway is to transform catechol to Krebs cycle intermediates (See > Chapter 4, Vol. 2, Part 2) (> Fig. 7). From phylogenetic analyses, proteobacterial catechol 1,2-dioxygenases (C12O) and actinobacterial C12Os form separate branches with low sequence identity to one another; however, all those enzymes seem to be characterized by similar kinetic properties and out of the chlorocatechols, only 4-chlorocatechol is transformed at a reasonable rate (Dorn and Knackmuss, 1978; Matsumura et al., 2004). Also proteobacterial and actinobacterial muconate cycloisomerases (MCI) constitute separate evolutionary branches with low sequence identity to one another (> Fig. 8).

. Figure 7 Diversity of pathways for the degradation/transformation of chlorocatechols. C12O, catechol 1,2-dioxygenase; CC12O, chlorocatechol 1,2-dioxygenase; MCI, muconate cycloisomerase; MCIP, proteobacterial muconate cycloisomerase; CMCIP, proteobacterial chloromuconate cycloisomerase; CMCIA, actinobacteral chloromuconate cycloisomerase; MLI, muconolactone isomerase; CMLIA, actinobacterial chloromuconolactone isomerase; DLHP, proteobacterial dienelactone hydrolase, DLHA, actinobacterial dienelactone hydrolase; tDLH, trans-dienelactone hydrolase; MAR, maleylacetate reductase. The unstable intermediate is enclosed in brackets.

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. Figure 8 Dendrogram showing the relatedness of muconate cycloisomerases (CatB) and chloromuconate cycloisomerases (ClcB, TfdD, TcbD, TetD). Mandelate racemase MdlA of P. putida is shown as outgroup.

Proteobacterial MCIs catalyze the cycloisomerization of 3-chloromuconate at reasonable rates (Ca´mara et al., 2007). This reaction is assumed to proceed via 4-chloromuconolactone as intermediate (> Fig. 7), which either spontaneously or enzyme-catalyzed rearranges with concomitant dehalogenation and decarboxylation to form protoanemonin (Blasco et al., 1995; Nikodem et al., 2003), a compound of high toxicity. The cycloisomerization product of 3-chloromuconate by actinobacterial MCIs has not yet been identified. In the case of 2-chloromuconate turnover, proteobacterial MCIs catalyze cycloisomerization only, to form both 2-chloro- and 5-chloromuconolactone as stable products (Vollmer et al., 1994) (> Fig. 7). Even though only few studies are available on actinobacterial MCIs, some studies suggested that they differ in catalytic properties from their proteobacterial counterparts and form exclusively 5-chloromuconolactone (Solyanikova et al., 1995). Both 5-chloro- and 2-chloromuconolactone are substrates of muconolactone isomerases of the 3-oxoadipate pathway. 5-Chloromuconolactone is transformed predominantly to cis-dienelactone probably via abstraction of the C4 proton followed by spontaneous chloride elimination (Prucha et al., 1996)

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and 2-chloromuconolactone into protoanemonin, probably by the elimination of CO2 and chloride from chlorosubstituted 3-oxoadipate enol-lactone (Skiba et al., 2002) (> Fig. 7). Mineralization of chlorocatechols via the 3-oxoadipate pathway is thus prevented by (1) restricted substrate specificity of the enzymes, (2) formation of protoanemonin as toxic dead-end product, or (3) formation of cis-dienelactone as dead-end product.

2.2.2

The Chlorocatechol Pathway

In most organisms isolated thus far, chlorocatechols are degraded by a chlorocatechol orthocleavage pathway (Reineke, 2001). This pathway is initiated by a broad specificity chlorocatechol 1,2-dioxygenase with consumption of molecular oxygen to produce the corresponding chloro-cis,cis-muconates (Dorn and Knackmuss, 1978). This reaction is identical to the one of the 3-oxoadipate pathway, and the main difference between the enzymes is their substrate specificity. A key enzyme in the pathway is chloromuconate cycloisomerase (CMCI), which differs in various aspects from muconate cycloisomerase. Proteobacterial CMCIs catalyze a specific cycloisomerization of 2-chloro-cis,cis-muconate (the metabolite of 3-chlorocatechol degradation) to form preferentially 5-chloromuconolactone, which is dehalogenated to transdienelactone (Vollmer and Schlo¨mann, 1995). Thus, proteobacterial CMCIs are specific dehalogenases. The formation of trans-dienelactone is due to the fact that, after cycloisomerization, the lactone ring has to rotate in the catalytic center to achieve proton abstraction and thus dehalogenation (Schell et al., 1999) (> Fig. 7). In contrast, chloride from the 3-position of 3-chloro-cis,cis-muconate (the metabolite of 4-chlorocatechol degradation) seems to be directly eliminated during cycloisomerization to form cis-dienelactone (Kaulmann et al., 2001) (> Fig. 7). Similarly, 2,4-dichloro-cis,cis-muconate, the metabolite of 3,5-dichlorocatechol degradation, is converted to 2-chloro-cis-dienelactone (Pieper et al., 1991). The dienelactones are converted into the respective maleylacetates by dienelactone hydrolases (DLH) (Schmidt and Knackmuss, 1980). Evidently, for the degradation of various chloroaromatics, an important prerequisite is that the dienelactone hydrolase is of relaxed substrate specificity and accepts both cis- and trans-isomers as substrates. The following enzyme, maleylacetate reductase, plays a major role in the degradation of chloroaromatic compounds either in the ortho-cleavage chlorocatechol pathway (Kaschabek and Reineke, 1992) or as part of the benzoquinol pathway. The original function is the reduction of the double bond to channel maleylacetate into the 3-oxoadipate pathway. Maleylacetates with chlorine substituents in the 2-position are subject to reductive dechlorination, e.g., 2-chloromaleylacetate, the intermediate in 3,5-dichlorocatechol degradation, is initially transformed to maleylacetate followed by reduction to 3-oxoadipate (Kaschabek and Reineke, 1992). Obviously, enzymatic attack on the C2-carbon results in an intermediate, which spontaneously eliminates chloride. Probably, dehalogenation of 2-chloromaleylacetate is a general capability of maleylacetate reductases. The ortho-cleavage chlorocatechol pathway described tolerates substitution at the aromatic ring of up to four chlorine atoms. Two dechlorination steps have been described up to now. The genes encoding the chlorocatechol pathway usually form clusters and the structures of the corresponding operons of chlorobenzoate degraders such as P. knackmussii B13 or C. necator NH9 or chlorobenzene degraders such as Pseudomonas sp. P51 or P. chlororaphis RW71 are nearly identical, in spite of the geographically distinct origins of the bacteria or the

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difference in their phylogenetic position (Pieper, 2005). Interestingly, the genetic organization of microorganisms isolated based on their capability to degrade 2,4-D usually differs substantially from that of chlorobenzoate and chlorobenzene degraders already described and in most strains the module comprises a gene encoding a phenol hydroxylase (tfdB), a protein probably involved in 2,4-D transport (tfdK) and a 2,4-D/a-ketoglutarate dioxygenase (tfdA). These gene clusters are typically missing tfdD and tfdF genes encoding chloromuconate cycloisomerases and maleylacetate reductases, respectively. However, in strains analyzed in this aspect, tfdD genes are located upstream of the tfdR regulator gene, whereas tfdF genes are located downstream of tfdA (Pieper, 2005). Even though analysis of chlorocatechol gene operons in proteobacteria had concentrated on b- and g-Proteobacteria, chlorocatechol genes have been also characterized also in members of the a-Proteobacteria (Mu¨ller et al., 2004). Both types of newly discovered chlorocatechol dioxygenases from Sphingomonas isolates represent deep-branching lines in the dendrogram and suggest a different reservoir and reduced transfer of the respective genes in a-Proteobacteria compared with the ones in b- and g-Proteobacteria. Only poor information is available on chlorocatechol genes in Actinobacteria, and thus far, only two modules from Rhodococcus opacus 1CP have been analyzed in detail (Maltseva et al., 1994a; Moiseeva et al., 2002). Usually, chlorocatechol 1,2-dioxygenases are characterized by their broad substrate specificity and can convert several higher chlorinated catechols. As an example, the chlorocatechol gene products of P. knackmussii B13 were capable of dealing with higher chlorinated catechols such as 3,5-dichloro- or 3,6-dichlorocatechol, whereas 3,4-dichlorocatechol was only a poor substrate (Dorn and Knackmuss, 1978; Oltmanns et al., 1988). In contrast, the enzyme of Pseudomonas sp. strain P51 was capable of transforming 3,4-dichlorocatechol (van der Meer et al., 1991). Recently, amino acids determining such differences in substrate specificity could be identified (Liu et al., 2005). In the case of chlorocatechol 1,2-dioxygenases from Rhodococcus, only one of the thus-far described enzymes exhibits broad substrate specificity (Moiseeva et al., 2002), whereas 4-chlorocatechol 1,2-dioxygenase prefers 4-chlorocatechol and has only negligible activity with 3-chlorocatechol (Maltseva et al., 1994a). All proteobacterial CMCIs obviously share common features, which discriminate them from the Rhodococcus enzymes, but also from MCIs. In agreement with their special biochemical properties, a phylogenetic comparison depicted proteobacterial CMCIs to constitute a separate subfamily, whereas the rhodococcal CMCIs, like the respective chlorocatechol 1,2-dioxygenases, appear to represent isolated evolutionary lines (> Fig. 8). As already described, proteobacterial CMCIs are specific dehalogenases, and capable of dehalogenating even higher substituted chloromuconates. In contrast, both CMCIs described from R. opacus 1CP exhibited extraordinary properties and catalyzed the formation of 5-chloromuconolactone from 2-chloromuconate but were not capable of dehalogenating (Moiseeva et al., 2002; Solyanikova et al., 1995) (> Fig. 7). Only some of the DLHs encoded in chlorocatechol gene clusters such as TfdEI from C. necator JMP134, and especially ClcD from P. knackmussii B13, have been described in more detail. These enzymes are active against both the cis- and trans-isomer, a feature that is indispensable for the mineralization of differently substituted chloroaromatics by proteobacteria as already described. Structure elucidation of the P. knackmussii B13 enzyme revealed that the protein belongs to the a/b hydrolase fold enzymes (Ollis and Nai, 1985). Analysis of

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DLHs from various chloroaromatic degraders showed that even though some of these DLHs share only low sequence identity (down to 13%), they all belong to the same family with a conserved Cys-His-Asp triad and highly conserved residues flanking the triad; however they all differ from enol-lactone hydrolases (Schlo¨mann, 1994). Unfortunately, only poor data are available on the substrate specificity of those DLHs; however, both DLHs from Rhodococcus are active only against the cis-isomer (Maltseva et al., 1994b; Moiseeva et al., 2002), contrasting DLHs from proteobacterial chlorobenzoate or chlorobenzene degraders.

2.2.3

The Alternative Pathway of 3-Chlorocatechol Degradation in Rhodococcus opacus 1CP

Among the Gram-positive organisms, chlorocatechol catabolism has mainly been investigated in R. opacus 1CP. 4-chloro- and 3,5-dichlorocatechol are transformed similar to the metabolism in proteobacteria, whereas 2-chloromuconate formed from 3-chlorocatechol is not dehalogenated during cycloisomerization. In contrast, dehalogenation is catalyzed by a muconolactone isomerase-related enzyme (> Fig. 7). Despite its sequence similarity to muconolactone isomerase, the Rhodococcus enzyme does not show muconolactone-isomerizing activity and thus represents an enzyme dedicated to its new function as a 5-chloromuconolactone dehalogenase forming cis-dienelactone (Moiseeva et al., 2002). Thus, a major difference between chlorocatechol degradation in proteobacteria and in Rhodococcus lies in the fact that cis-dienelactone is the intermediate of both 4-chloro- and 3-chlorocatechol degradation in Rhodococcus, whereas cis- and trans-dienelactone, respectively, are formed by proteobacteria, therefore requiring a DLH with broad substrate specificity (> Fig. 7).

2.2.4

The Alternative Pathway of 4-Halocatechol Degradation

Another variant of the chlorocatechol pathway has been described in Pseudomonas reinekei MT1 (Nikodem et al., 2003). In this strain 4-chlorocatechol formed from 4-chloro- and 5-chlorosalicylate is channeled into the 3-oxoadipate pathway by a catechol 1,2-dioxygenase and a MCI that showed unusual kinetic properties as being adapted for turnover of 4-chlorocatechol and 3-chloromuconate, respectively (Ca´mara et al., 2007). A trans-dienelactone hydrolase capable of transforming the trans- but not the cis-dienelactone isomer has been observed to be crucial for 4-chlorocatechol degradation by this strain. It was shown that the enzyme interacts with the MCI-catalyzed transformation of 3-chloromuconate and hydrolyzes the cycloisomerization intermediate 4-chloromuconolactone, thereby preventing the formation of protoanemonin in favor of maleylacetate (Nikodem et al., 2003) (> Fig. 7). Maleylacetate is then transformed by maleylacetate reductase. DLHs active against trans-dienelactone only and not able to transform cis-dienelactone, the intermediate of 4-chlorocatechol degradation by enzymes of the chlorocatechol pathway, have initially been described to be involved in 4-fluorocatechol degradation by C. necator 335 and C. necator JMP222 and probably responsible for further degradation of intermediate 4-fluoromuconolactone (Schlo¨mann et al., 1990).

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Chlorocatechol Degradation via the Meta-Cleavage Pathway

Catechol meta-cleavage routes are widespread and usually involved in the degradation of methylsubstituted compounds such as toluene or methylphenols (See > Chapter 4, Vol. 2, Part 2). For a long time the presence of such a meta-cleavage pathway was assumed to severely interfere with the degradation of chloroaromatics. One of the reasons of interference was assumed to be the formation of a reactive acyl chloride, e.g., from 3-chlorocatechol by the catechol 2,3-dioxygenase of P. putida mt-2 (XylE) (Bartels et al., 1984), resulting in irreversible inactivation of the ring cleavage enzyme (> Fig. 9). In other cases, reversible inactivation was shown to be due to a rapid oxidation of the active site ferrous iron to its ferric form with concomitant loss of activity (Vaillancourt et al., 2002), and a general mechanism for the inactivation of extradiol dioxygenases during catalytic turnover involving the dissociation of superoxide from the enzyme-catechol-dioxygen ternary complex was suggested. In contrast to 3chlorocatechol, 4-chlorocatechol is a moderate substrate for various catechol 2,3-dioxygenases (Bartels et al., 1984; Murray et al., 1972), among them catechol 2,3-dioxygenases of family I.2.A (Eltis and Bolin, 1996), which have been commonly observed to be involved in the degradation of methylaromatics. However, despite high sequence identity, members of this subfamily exhibit very different substrate specificities, and the capability to transform 4-chlorocatechol is not a general characteristic of catechol 2,3-dioxygenases of family I.2.A (Kitayama et al., 1996). Some publications postulate that compounds degraded via catechols chlorinated in the 4-position might be mineralized via a catechol degrading meta-cleavage pathway (Arensdorf and Focht, 1995) but information about the way in which the products are dechlorinated is missing. In 1997, the first isolate, P. putida GJ31, capable of degrading a chloroaromatic compound (chlorobenzene) via 3-chlorocatechol using a meta-cleavage pathway was described (Mars et al., 1997). The CbzE chlorocatechol 2,3-dioxygenase of strain GJ31 productively converts 3-chlorocatechol, and stoichiometric displacement of chloride leads to the production of 2-hydroxymuconate (Kaschabek et al., 1998), which is further converted through the metacleavage pathway (> Fig. 9). 3-Chlorocatechol was found to be the preferred substrate of CbzE. Additional pseudomonads using a meta-cleavage route for 3-chlorocatechol degradation were isolated in the meantime from various contaminated environments (Go¨bel et al., 2004), suggesting that productive meta-cleavage of 3-chlorocatechol is not atypical for chloroaromatic degradation. All the three isolates harbor chlorocatechol 2,3-dioxygenases, highly resistant to inactivation during 3-chlorocatechol turnover, sharing 97% amino acid sequence identity with the enzyme from strain GJ31, thus forming a distinct subfamily of catechol 2,3dioxygenases in what was termed family I.2.C. Other extradiol dioxygenases of family 1.2.C do not share the capability of effectively transforming 3-chlorocatechol. In all the chlorobenzene-degrading strains already mentioned, cbzT genes encoding ferredoxins are located upstream of cbzE. Similar ferredoxins have been observed to be encoded in various meta-cleavage pathways (Hugo et al., 2000). Catechol 2,3-dioxygenases are oxygen sensitive and unstable in vitro, particularly in the presence of substituted catechol substrates. This instability was shown to be due to the oxidation of active site Fe(II)–Fe(III) (Vaillancourt et al., 2002), and CbzT-like ferredoxins reactivate catechol 2,3-dioxygenases through the reduction of the iron atom in the active site of the enzyme (Hugo et al., 2000). It was thus proposed that the ability of strain GJ31 to metabolize both chlorobenzene and toluene might depend on the regeneration of the chlorocatechol dioxygenase activity mediated by CbzT (Tropel et al., 2002).

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. Figure 9 Meta-cleavage pathways for catechol, 3-chlorocatechol, protocatechuate and 5-chloroprotocatechuate. The acylchlorides are shown in brackets. The 3-chlorocatechol ring-cleavage product can react either with the ring cleavage enzyme resulting in suicide inactivation, or be rapidly hydrolyzed to give 2-hydroxymuconate. The 5-chloroprotocatechuate ring-cleavage product undergoes intramolecular rearrangement and dehalogenation.

2.2.6

Degradation of 5-Chloroprotocatechuate via a Meta-Cleavage Pathway

A productive meta-cleavage without suicide effect has been known for many years. The extradiol cleavage of 5-chloroprotocatechuate by protocatechuate 4,5-dioxygenase (Kersten et al., 1985) results in the formation of 2-pyrone-4,6-dicarboxylate by nucleophilic displacement of chloride (> Fig. 9). This indicates that cyclization entailing nucleophilic displacement of halogen provides an effective alternative to the enzyme suicide inactivation that occurs when a nucleophilic group of the extradiol dioxygenase undergoes acylation. An important aspect of this mechanism is that the ring fission product remains bound to the enzyme during the complete configuration change that precedes nucleophilic displacement.

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2-Pyrone-4,6-dicarboxylate is a metabolite of the protocatechuate 4,5-dioxygenase pathway typically formed after the dehydrogenation of the ring-cleavage product (see > Fig. 9), such that both protocatechuate and 5-chloroprotocatechuate degradation merge at this metabolic intermediate. So far, only the mineralization of 3-chlorobenzoate by C. testosteroni BR60 (Nakatsu et al., 1997) has been proposed to occur via such a route.

2.3

Ring Cleavage of (Chloro)Benzoquinols and (Chloro) Hydroxybenzoquinols

Ring cleavage dioxygenases involved in the turnover of (chloro) benzoquinols and (chloro) hydroxybenzoquinols has been identified from various microorganisms degrading g-HCH, or chlorophenols (> Fig. 10). Chlorobenzoquinol (but also benzoquinol) produced during the catabolism of g-HCH in S. japonicum UT26 is subject to direct ring cleavage by a novel type of extradiol dioxygenase, designated chlorobenzoquinol/benzoquinol 1,2-dioxygenase (LinE), which preferentially cleaves aromatic rings with two hydroxyl groups at para positions (Miyauchi et al., 1999) and produces maleylacetate from either chlorobenzoquinol or benzoquinol (> Fig. 10). Highly similar or identical dioxygenases (>99% sequence identity) have been identified in other g-HCH-degrading organisms (Dogra et al., 2004). More distantly related extradiol dioxygenases (51–53% sequence identity) and designated 2,6-dichlorobenzoquinol 1,2-dioxygenases (PcpA) were observed in PCP degrading strains such as S. chlorophenolicum ATCC 39723 (Xu et al., 1999) and produce 2-chloromaleylacetate (2-CMA) from 2,6-dichlorobenzoquinol (> Fig. 10). A protein highly similar to PcpA proteins and with activity with 2,6-dichlorobenzoquinol was also observed in S. japonicum UT26 (LinEb exhibiting 92% sequence identity to PcpA proteins (Endo et al., 2005)). However, its involvement in g-HCH degradation responsible for cleavage of 2,5-dichlorobenzoquinol is unlikely, because such a cleavage would result in the formation of 3-chloromaleylacetate, which cannot be dehalogenated by maleylacetate reductase. In contrast, members of the intradiol dioxygenase family are involved in the degradation of hydroxybenzoquinols formed from trichlorophenols, 2,4-dichloro- or 4-chlorophenol, and these (chloro)hydroxybenzoquinol dioxygenases form a defined phylogenetic group inside the intradiol dioxygenase family (See > Chapter 39, Vol. 2, Part 6). Hydroxybenzoquinol is cleaved by TftH hydroxybenzoquinol 1,2-dioxygenase to yield maleylacetate during the mineralization of 2,4,5-T by B. phenoliruptrix AC1100 (Daubaras et al., 1996) (> Fig. 10). In 2,4,6-TCP degradation by C. necator JMP134, TcpC 6-chlorohydroxyquinol 1,2-dioxygenase is responsible for direct cleavage of 6-chlorohydroxybenzoquinol producing 2-chloromaleylacetate (Louie et al., 2002) (> Fig. 10). An important difference between the pathways of 2,4,5TCP and 2,4,6-TCP degradation is the fact that 6-chlorohydroxybenzoquinol is directly cleaved instead of a preceding reductive dechlorination forming hydroxybenzoquinol. Interestingly, TftH hydroxybenzoquinol 1,2-dioxygenase from B. phenoliruptrix AC1100, which shares 53% amino acid sequence identity with TcpC, is unable to use 5-chloro- or 6chlorohydroxybenzoquinol (Daubaras et al., 1996). The catabolism of (chloro)benzoquinols and (chloro)hydroxybenzoquinols is completed by the conversion of MA or 2-CMA to 3-oxoadipate (> Fig. 10) by the action of maleylacetate reductases as already described (see Section 2.2.2).

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. Figure 10 Catabolic pathways for (chloro)benzoquinols and (chloro)hydroxybenzoquinols. Unstable intermediates are enclosed in brackets. Broken arrows indicate reactions of the upper pathway detailed in > Fig. 6.

3

Research Needs

Most of the current knowledge on biochemistry and genetics of chloroaromatic aerobic microbial metabolism has been obtained with bacterial strains isolated by traditional culture-dependent approaches. This is a severe drawback to understanding microbial degradation of chloroaromatics, as only a very minor proportion of microbes inhabiting the different ecosystems can be isolated by standard procedures and only a few reports are available on microbial associations (consortia) degrading this kind of compounds. Although it is possible that current knowledge covers a significant proportion of microbial strategies to degrade chloroaromatics, it would also be possible that new, unexpected ways to metabolize these compounds remain still unknown in the significant fraction of yet-uncultured microbes. Functional metagenomics approaches are required to address this point. Since natural haloaromatic formation is increasingly evident, studies including diverse ecosystems, not

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necessarily exposed to chloroaromatic pollutants, will also provide a better idea on the diversity of metabolic pathways for aerobic degradation of chloroaromatics. Several cases of adaptation or recruitment of pathways involved in the metabolism of natural aromatics for metabolism of the corresponding chloroaromatic counterparts are already reported. A further exploration of these possibilities would provide new insights into these phenomena. For instance, degradation of aromatic compounds through CoA derivatives seems to play a more significant role in aerobic degradation than initially expected (See > Chapter 4, Vol. 2, Part 2). CoA derivatives have been reported as intermediates of 4-chlorobenzoate degradation, but additional routes are expected to be discovered. Available are only a few studies of the degradation of mixed substituted chloroaromatic compounds, i.e., carrying additional amino/nitro/sulfonic substituents or heterocyclic aromatics, which without doubt, may give interesting new insights into aerobic microbial degradation of pollutants. Even for the pathways already reported, there are several unclarified questions that deserve attention. For example, by which mechanism is the third or fourth chlorosubstituent eliminated during trichloro- or tetrachlorocatechol metabolism? What are the biochemical grounds for the degradation of 4-chlorocatechol via the meta-cleavage? By which mechanism is 3,6-dichlorogentisate mineralized in dicamba degrading microorganisms?

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6 Aerobic Degradation of Halogenated Aliphatics S. Fetzner Institut fu¨r Molekulare Mikrobiologie und Biotechnologie, Westfa¨lische Wilhelms-Universita¨t Mu¨nster, Mu¨nster, Germany [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 866 2 Halomethanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 866 3 Halo-n-Alkanes and Haloalkanoic Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 873 4 Haloalkenes and Haloalkenoic Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 876 5 Haloalcohols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 878 6 Halogenated Alicyclic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 880 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 881

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_62, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Many mono- and dihalogenated aliphatic compounds support bacterial growth, whereas highly halogenated substances in the majority of cases are recalcitrant to aerobic biodegradation. Pathways for the bacterial degradation of halogenated methane, n-alkanes, alkanoic acids, alkenes, alkenoic acids, alcohols, and the alicyclic insecticide Lindane have been elucidated and are briefly summarized in this chapter. Dehalogenases, which catalyze the cleavage of the carbon-halogen bond, are the key enzymes of these catabolic pathways. They use different catalytic mechanisms and belong to different protein families, reflecting their diverse evolutionary origin. Enantioselective dehalogenases, especially halohydrin dehalogenases, are attractive for biocatalytic applications.

1

Introduction

Halogenated organic compounds are among industrially important solvents, building blocks for chemical synthesis, and pesticides. Many of these synthetic compounds, especially the chlorinated insecticides, were intended to persist in the environment and hence emerged as environmental pollutants. Factors in their recalcitrance to biodegradation may include poor bioavailability (e.g., due to poor solubility, or strong sorption to soil), a xenobiotic structure (e.g., a high degree of halogen substitution) which is not recognized by existing uptake proteins and catabolic enzymes, or high toxicity of the compound itself or of intermediates which accumulate in incomplete catabolic pathways. With only a few exceptions, those haloorganics that constitute the bulk quantities in the biosphere are synthesized industrially (Anonymous, 2008; Fetzner, 1998). On the other hand, thousands of halogenated compounds are produced naturally by a variety of bacteria, algae, fungi, and higher eukaryotes, and are also formed during natural abiogenic processes (Gribble, 2003), substantiating that the carbon-halogen bond per se is not xenobiotic. This chapter gives a brief overview of pathways of aerobic bacterial degradation of halogenated aliphatics, and introduces the enzymes which catalyze the critical dehalogenation step. > Table 1 specifies a number of bacterial strains which are able to utilize haloaliphatic compounds as growth substrates, and > Table 2 gives examples of dehalogenases active towards haloaliphatic compounds, with a focus on their substrate specificities.

2

Halomethanes

Chloromethane (methyl chloride), the most abundant halocarbon in the atmosphere, is a naturally occurring metabolite produced mainly by marine algae and wood-rotting fungi, and generated by biomass combustion (McDonald et al., 2002; Scha¨fer et al., 2007). Bromomethane is also emitted by natural processes, but use of synthetic bromomethane as a soil fumigant in agriculture significantly contributed to annual emissions, until its use was curtailed by the ‘‘Montreal Protocol on Substances That Deplete the Ozone Layer’’ (UNEP, 2006). Cometabolic conversion of chloromethane by aerobic bacteria includes hydrolytic reactions as well as fortuitous oxidation to formaldehyde by methanotrophic and nitrifying bacteria which produce non-specific methane monooxygenase and ammonia monooxygenase,

Aerobic Degradation of Halogenated Aliphatics

6

. Table 1 Examples of halogenated aliphatics utilized as growth substrates by aerobic bacteria Compound

Organism

Reference

Halomethanes Chloromethane

Hyphomicrobium sp. MC1

Hartmans et al. (1986)

Hyphomicrobium spp.

McAnulla et al. (2001a)

Hyphomicrobium chloromethanicum CM2

McAnulla et al. (2001b)

Methylobacterium chloromethanicum CM4

McDonald et al. (2001)

Nocardioides sp. SAC-4

McAnulla et al. (2001a)

Chloromethane and bromomethane

Aminobacter lissarensis CC495 Coulter et al. (1999)

Chloro-, bromo-, iodomethane

Aminobacter ciceronei IMB1

Connell Hancock et al. (1998)

Leisingera methylohalidivorans Schaefer et al. (2002) MB2 Pseudomonas sp. DM1

Brunner et al. (1980)

Hyphomicrobium sp. DM2

Stucki et al. (1981)

Methylobacterium dichloromethanicum DM4

Ga¨lli and Leisinger (1985)

Methylophilus sp. DM11

Scholtz et al. (1988)

Bromoethane

Acinetobacter sp. GJ70

Janssen et al. (1987)

1,2-Dichloroethane

Xanthobacter autotrophicus GJ10

Janssen et al. (1985)

Ancylobacter aquaticus AD20, AD25, AD27

van den Wijngaard et al. (1992)

1,2-Dibromoethane

Mycobacterium sp. GP1

Poelarends et al. (1999)

1-Bromopropane

Mycobacterium sp. GP1

Poelarends et al. (1999)

1-Chloropropane

Xanthobacter autotrophicus GJ10

Janssen et al. (1985)

Mycobacterium sp. GP1

Poelarends et al. (1999)

1,3-Dichloropropane

Xanthobacter autotrophicus GJ10

Janssen et al. (1985)

1-Chlorobutane

Xanthobacter autotrophicus GJ10

Janssen et al. (1985)

Rhodococcus sp. m15–3 (formerly, Corynebacterium sp. m15–3)

Yokota et al. (1987)

Dichloromethane

Halo-n-Alkanes

Acinetobacter sp. GJ70

Janssen et al. (1987)

1-Chloropentane

Acinetobacter sp. GJ70

Janssen et al. (1987)

C2–C8 1-haloalkanes

Arthrobacter sp. HA1

Scholtz et al. (1987)

867

868

6

Aerobic Degradation of Halogenated Aliphatics

. Table 1 (Continued) Compound

Organism

Reference

C3,C4,C5,C6,C12,C14,C16,C18 1-chloroalkanes

Rhodococcus erythropolis Y2

Armfield et al. (1995)

C3–C18 1-chloroalkanes

Rhodococcus rhodochrous NCIMB 13064

Curragh et al. (1994) Belkin (1992)

C6–C18 1-bromoalkane

Pseudomonas sp. ES-2

C6 and C9 a,o-dichloroalkane

Acinetobacter sp. GJ70

Janssen et al. (1987)

1,10-dichlorodecane

Pseudomonas sp. 273

Wischnak et al. (1998)

Burkholderia (formerly, Pseudomonas) cepacia MBA4

Tsang et al. (1988)

Haloalkanoic acids Chloroacetate

Alcaligenes sp. CC1

Kohler-Staub and Kohler (1989)

Ancylobacter aquaticus AD20, AD25, AD27

van den Wijngaard et al. (1992)

Alcaligenes xylosoxidans ABIV

Brokamp and Schmidt (1991)

Bromoacetate

Burkholderia cepacia MBA4

Tsang et al. (1988)

2-Chloropropionate

Pseudomonas putida PP3

Slater et al. (1979)

Pseudomonas sp. 113

Motosugi et al. (1982a)

Pseudomonas sp. YL

Liu et al. (1994)

Alcaligenes sp. CC1

Kohler-Staub and Kohler (1989)

Ancylobacter aquaticus AD20, AD25, AD27

van den Wijngaard et al. (1992)

Alcaligenes xylosoxidans ABIV

Brokamp and Schmidt (1991)

Pseudomonas putida PP3

Slater et al. (1979)

Burkholderia cepacia MBA4

Tsang et al. (1988)

Pseudomonas putida PP3

Slater et al. (1979)

Alcaligenes xylosoxidans ABIV

Brokamp and Schmidt (1991)

2-Bromopropionate 2,2-Dichloropropionate

Bradyrhizobium sp. DA2, DA3, Marchesi and Weightman DA4 (2003)

2- and 3-Chlorobutyrate

Brucella strain DA5

Marchesi and Weightman (2003)

Alcaligenes sp. CC1

Kohler-Staub and Kohler (1989)

Mycobacterium aurum L1

Hartmans and de Bont (1992)

Haloalkenes Chloroethene (Vinyl chloride)

1,3-Dichloropropene

Mycobacterium spp.

Coleman and Spain (2003)

Nocardioides sp. JS614

Mattes et al. (2005)

Pseudomonas pavonaceae 170 Poelarends et al. (1998) (formerly, P. cichorii 170)

Haloalkenoic acids 2-Chloroacrylate

Pseudomonas sp. YL

Liu et al. (1994)

Aerobic Degradation of Halogenated Aliphatics

6

. Table 1 (Continued) Compound 3-Chloroacrylate

Organism

Reference

Pseudomonas cepacia CAA1

Hartmans et al. (1991)

Coryneform strain CAA2

Hartmans et al. (1991)

Coryneform strain FG41

van Hylckama Vlieg and Janssen (1992)

Pseudomonas pavonaceae 170 Poelarends et al. (1998) 3-Chlorocrotonate

Alcaligenes sp. CC1

Kohler-Staub and Kohler (1989)

Pseudomonas sp. CE1r

Stucki and Leisinger (1983)

Haloalcohols 2-Chloroethanol

Pseudomonas putida US2

Strotmann et al. (1990)

2,3-Dichloro-1-propanol

Pseudomonas sp. OS-K-29

Kasai et al. (1990)

2,3-Dibromo-1-propanol

Flavobacterium sp.

Castro and Bartnicki (1968)

1,3-Dichloro-2-propanol

Agrobacterium radiobacter AD1

van den Wijngaard et al. (1989)

Arthrobacter sp. AD2

van den Wijngaard et al. (1989)

Agrobacterium radiobacter AD1

van den Wijngaard et al. (1989)

Arthrobacter sp. AD2

van den Wijngaard et al. (1989)

3-Chloro-1,2-propanediol

2-Chloroallylalcohol 3-Chloroallylalcohol

Pseudomonas sp. JD1 and JD2 van der Waarde et al. (1993) Pseudomonas sp.

Belser and Castro (1971)

Pseudomonas pavonaceae 170 Poelarends et al. (1998) Halogenated alicyclic compounds g-Hexachlorocyclohexane (Lindane)

Sphingobium japonicum (formerly, Sphingomonas paucimobilis) UT26

Imai et al. (1989), Nagata et al. (2007)

respectively. However, a number of aerobic methylotrophic bacteria are capable of growth on chloro- and bromomethane (Scha¨fer et al., 2007). The metabolism of chloromethane utilization has been studied most extensively in Methylobacterium chloromethanicum CM4 by Leisinger and colleagues (Vannelli et al., 1999; reviewed by McDonald et al., 2002 and Scha¨fer et al., 2007). The first step of the pathway, catalyzed by the CmuA protein which contains a methyltransferase domain and a corrinoid-binding domain, involves transfer of the methyl group of chloromethane to the corrinoid group of CmuA. The methyl group is then passed to tetrahydrofolate and oxidized to formate and finally to CO2, providing reducing equivalents for biosyntheses (> Fig. 1). Based on biochemical data and sequence analysis of cmu gene clusters, it was suggested that a similar pathway is operative in Hyphomicrobium chloromethanicum CM2, Aminobacter lissarensis CC495, and Aminobacter ciceronei IMB-1. However, the CmuA pathway is possibly not the only bacterial halomethane utilization pathway (Scha¨fer et al., 2007). Dichloromethane (methylene chloride), which is produced by chemical synthesis for use as an industrial solvent, is oxidized by Pseudomonas, Hyphomicrobium, Methylobacterium and Methylophilus strains. The dichloromethane-utilizing methylotrophs possess a dehalogenase

869

Xanthobacter autotrophicus GJ10

Sphingobium japonicum (formerly, Sphingomonas paucimobilis) UT26

a/b-Hydrolase fold C1–C6 1-chloro-n-alkanes, C1–C5 1-bromo-n-alkanes, 1-iodomethane, dichloromethane, 1-iodopropane, C2, C3, C6, C9, a,o-dichloro-n-alkanes, 1,2-dibromoethane, 1,2-dichloropropane, 1,2-dibromopropane, 6-chlorohexanol, 6-bromohexanol, epichlorohydrin, epibromohydrin, 3-chloropropene

C3–C14 1-chloroalkanes, C2, C4 1-bromoalkanes, C3,C4, a/b-Hydrolase fold C6 1-iodoalkanes, 2-chlorobutane, 2-chlorooctane, 3-chlorohexane, C3–C10 a,o-dichloro-n-alkanes, 1,2-dibromoethane, 1-bromo-2-chloroethane, 1,1-dichloroethane, 1,1,1-trichloroethane, 1,2-dibromopropane, 1,3-dibromopropane, 1,3-diiodopropane, 1-bromo-3-chloropropane, 2-bromo-1-chloropropane, 1-bromo2-methylpropane, (1,2,3-trichloropropane), C3,C4,C8 2-chloro-n-alkane, 3-chlorohexane, bis(2-chloroethyl) ether, 2-chloroethanol, 2-bromoethanol, 4-chlorobutanol, 6-chlorohexanol, 6-bromohexanol, chloroacetate, 2-chlorobutyrate, bromocyclohexane, chlorocyclohexane, bromocycloheptane, bromomethylcyclohexane, 3-chloropropene, 3-chloro-2-methylpropene, 3-chloro-2-(chloromethyl)1-propene, 3,4-dichlorobutene, cis- and trans-1,4-dichloro-2-butene, 2,3-dichloropropene, (cis/trans)-1,3-dichloropropene, 4-bromobutyronitril

LinB, haloalkane dehalogenase (1,3,4,6tetrachloro-1,4cyclohexadiene hydrolase)

Hyphomicrobium sp. DM2

Organism

DhlA, haloalkane dehalogenase

Glutathione S-transferase (GST)

Protein (super) family

Dichloromethane, chlorobromomethane, dibromomethane, diiodomethane

Substrates

Nagata et al. (1997), Marek et al. (2000), Damborsky´ et al. (2001), Prokop et al. (2003), Oakley et al. (2004), Chaloupkova´ et al. 2003, Kmunı´cek et al. (2005), Monincova´ et al. (2007)

Keuning et al. (1985), Verschueren et al. (1993), Damborsky´ et al. (2001)

Kohler-Staub and Leisinger (1985)

References

6

DcmA, dichloromethane dehalogenase

Enzyme

. Table 2 Examples of dehalogenases active towards haloaliphatic compounds

870 Aerobic Degradation of Halogenated Aliphatics

monochloroacetate, monobromoacetate, monoiodoacetate, dichloroacetate, trichloroacetate, D-2-chloropropionate, L-2-chloropropionate, DL-2-bromopropionate, 2,2-dichloropropionate, DL-2-chloro-n-butyrate, DL-2-bromo-n-butyrate, DL-2-bromo-n-valerate

monochloroacetate, monobromoacetate, dichloroacetate, dibromoacetate, DL-2-chloropropionate, 2,3-dichloropropionate, DL-2-chlorobutyrate

monochloroacetate, monobromoacetate, monoiodoacetate, L-2-chloropropionate, 2,2-dichloropropionate, DL-2-chloro-n-butyrate, DL-2-bromo-n-valerate

1,3-dichloro-2-propanol, 1-chloro-2-propanol, 3-chloro-1,2-propanediol, 2-chloroethanol, 2-bromoethanol, 1-bromo-2-propanol, 1,3dibromo-2-propanol, 2-bromoethanol, 1,3dichloroacetone, chloroacetone, (R)-2-chloro-1phenylethanol, (S)-2-chloro-1-phenylethanol

1,3-dichloro-2-propanol, (R)-2,3-dichloro-1-propanol, 3-chloro-1,2-propanediol, 2-chloroethanol, 2-bromoethanol, 1,3-dibromo-2-propanol, chloroacetone, 2-phenyl-1-chloro-2-ethanol, (R)2-chloro-1-phenylethanol, [(S)-2-chloro-1phenylethanol], (R)-p-nitro-2-bromo-1-phenylethanol

trans-3-chloroacrylate, trans-3-bromoacrylate, 2-oxo-3-pentynoate (hydration)

DL-DEX, DL-2-haloacid dehalogenase

DhlB, L-2-haloacid dehalogenase

L-DEX, L-2-haloacid dehalogenase

HheA, halohydrin dehalogenase

HheC, halohydrin dehalogenase

CaaD, trans-3-chloroacrylic acid dehalogenase

Lutje Spelberg et al. (1999), van Hylckama Vlieg et al. (2001), de Jong et al. (2003, 2005), Tang et al. (2003)

Agrobacterium radiobacter AD1

SDR

4-Oxalocrotonate tautomerase

Poelarends et al. (2001), Wang et al. (2003), de Jong et al. (2004)

van den Wijngaard et al. (1991), van Hylckama Vlieg et al. (2001), de Jong et al. (2006)

Arthrobacter sp. AD2

Short-chain dehydrogenase reductase (SDR)

Pseudomonas pavonaceae 170

Liu et al. (1994), Hisano et al. (1996), Li et al. (1998)

Pseudomonas sp. YL

Group II haloacid dehalogenase (HAD)

Group II haloacid dehalogenase (HAD)

van der Ploeg et al. (1991), Ridder et al. (1999)

Pseudomonas sp. 113 Motosugi et al. (1982b), Nardi-Dei et al. (1999)

Xanthobacter autotrophicus GJ10

Group I haloacid dehalogenase

Aerobic Degradation of Halogenated Aliphatics

6 871

Substrates

LinA, hexachlorocyclohexane (HCH) dehydrochlorinase

New family within tautomerase superfamily

Protein (super) family

a-hexachlorocyclohexane (HCH), g-HCH, d-HCH, Dehydratase a-pentachlorocyclohexene, g-pentachlorocyclohexene

cis-CaaD, cis-3-chloroacrylic cis-3-chloroacrylate, cis-3-bromoacrylate, 2-oxo-3acid dehalogenase pentynoate (hydration)

Enzyme

References

Sphingobium japonicum UT26

Nagata et al. (1993, 1997)

Coryneform Poelarends et al. bacterium strain FG41 (2004), de Jong et al. (2007)

Organism

6

. Table 2 (Continued)

872 Aerobic Degradation of Halogenated Aliphatics

Aerobic Degradation of Halogenated Aliphatics

6

. Figure 1 Proposed pathway for chloromethane metabolism in Methylobacterium chloromethanicum CM4 (modified from McDonald et al., 2002; Vannelli et al., 1999). CmuA, methyltransferase I (corrinoid protein); CmuB, methyltransferase II; MetF, putative 5,10-methylene-tetrahydrofolate reductase/ 5,10-methenyl-tetrahydrofolate cyclohydrolase; PurU, putative 10-formyl-tetrahydrofolate hydrolase; FDH, formate dehydrogenase.

. Figure 2 Reaction catalyzed by dichloromethane dehalogenase (DcmA). GSH, glutathione.

(DcmA) which requires glutathione (GSH) as cosubstrate. Chloride is displaced in a nucleophilic substitution reaction, yielding S-chloromethyl glutathione which decomposes to chloride, GSH, and formaldehyde, a central metabolite of methylotrophic growth (> Fig. 2) (Leisinger et al., 1994). Analysis of the organization of the dcm regions of a number of dichloromethane-utilizing strains revealed the presence of IS (insertion sequence) elements and suggested that a transposon is involved in horizontal transfer of the gene regions involved in dichloromethane metabolism (Schmid-Appert et al., 1997). In contrast to mono- and dichloromethane, which are utilized as growth substrates, trichloromethane (chloroform) and tetrachloromethane (carbon tetrachloride) are not productively metabolized by aerobic bacteria, however, chloroform is co-metabolically mineralized to CO2 by bacteria producing methane monooxygenase or toluene monooxygenases (Jahng and Wood, 1994; McClay et al., 1996). Tetrachloromethane is only amenable to anaerobic transformations.

3

Halo-n-Alkanes and Haloalkanoic Acids

Chloroalkanes are widely used as plasticizers, solvents, lubricants, and fire retardants. 1,2-Dichloroethane (ethylene dichloride) is manufactured in huge quantities, primarily for use in the synthesis of vinyl chloride. In 2007, for example, about 16 million metric tons of 1,2-dichloroethane were produced in the US, Europe, and Japan (Anonymous, 2008).

873

874

6

Aerobic Degradation of Halogenated Aliphatics

The bacterial degradation of haloalkanes can proceed through different pathways: Whereas short-chain (C2–C6) 1-halo-n-alkanes and a,o-dihalo-n-alkanes are usually hydrolytically dechlorinated to form the corresponding alcohols, which are further oxidized to fatty acids, dechlorination of long-chain mono- and a,o-disubstituted haloalkanes seems to require oxygenase-type enzymes. Growth of Xanthobacter autotrophicus GJ10 and Ancylobacter aquaticus strains on 1,2dichloroethane requires two hydrolytic dehalogenases, haloalkane dehalogenase DhlA, and haloacid dehalogenase DhlB (> Fig. 3a) (for reviews, see: de Jong and Dijkstra, 2003;

. Figure 3 Degradation of 1,2-dihaloalkanes (modified from van Hylckama Vlieg et al., 2000). (a) Degradation of 1,2-dichloroethane by Xanthobacter autotrophicus GJ10. (1) 1,2-Dichloroethane; (2) 2-chloroethanol; (3) chloroacetaldehyde; (4) chloroacetic acid; (5) glycolate; DhlA, haloalkane dehalogenase; Mox, alcohol dehydrogenase; Ald, aldehyde dehydrogenase; DhlB, 2-haloacid dehalogenase. (b) Degradation of 1,2-dibromoethane by Mycobacterium sp. GP1. (1) 1,2-Dibromoethane; (2) 2-bromoethanol; (3) epoxyethane; DhaA, haloalkane dehalogenase; HheB, halohydrin dehalogenase.

Aerobic Degradation of Halogenated Aliphatics

6

Pries et al., 1994; van Hylckama Vlieg et al., 2000). However, bacteria that grow on haloalkanes or 1,2-dichloroethane are usually inhibited by 1,2-dibromoethane, a compound that has been used as pesticide and insecticidal fumigant. This is caused by the higher reactivity of bromoacetaldehyde compared with chloroacetaldehyde, resulting in inactivation of the aldehyde dehydrogenase (van Hylckama Vlieg et al., 2000). Interestingly, Mycobacterium sp. strain GP1, which utilizes 1,2-dibromoethane as sole carbon source, has evolved a pathway that avoids formation of the toxic bromoacetaldehyde, but after initial hydrolytic debromination by haloalkane dehalogenase uses a halohydrin dehalogenase to form epoxyethane from 2-bromoethanol (> Fig. 3b) (Poelarends et al., 1999). Halogenated propanes, such as 1,2,3-tribromopropane and 1,2-dibromo-3-chloropropane, were formerly applied as nematocides in agriculture; 1,2,3-trichloropropane is used as intermediate in organic syntheses and as a solvent. However, aerobic soil bacteria with the ability to grow on trihalogenated propanes have not been described. To construct an organism that degrades di- and tribromopropanes, Bosma et al. (1999) expressed the gene encoding haloalkane dehalogenase DhaA from Rhodococcus sp. m15–3 in Agrobacterium radiobacter AD1, which has the ability to grow on dihalogenated propanols. The recombinant strain indeed utilizes both 1,2,3-tribromopropane and 1,2-dibromo-3-chloropropane as carbon sources. Improving the activity of DhaA towards 1,2,3-trichloropropane in a directed evolution approach and expression of the mutated dhaA gene in A. radiobacter AD1 moreover resulted in a recombinant strain growing on 1,2,3-trichloropropane (Bosma et al., 2002). Haloalkane dehalogenases (haloalkane halidohydrolases), which catalyze the initial key step of the bacterial degradation of short-chain haloalkanes, like many other hydrolytic enzymes belong to the a/b-hydrolase fold family. X-ray structures of three haloalkane dehalogenases (DhlA, DhaA, and LinB) have been solved, and a wealth of biochemical and structural data suggests that they share a similar catalytic mechanism (reviewed in: de Jong and Dijkstra, 2003; Janssen, 2004). Five key residues are essential for catalysis: Three residues (Asp-His-Asp/Glu), which are functionally similar to the ‘‘classical’’ Ser-His-Asp/Glu catalytic triad of serine hydrolases, are involved in catalysis. The nucleophilic aspartate attacks the haloalkane substrate to form a covalent Asp-substrate intermediate with concomitant release of halide, and subsequently the cosubstrate H2O, activated by the His-Asp/Glu pair, hydrolyzes the covalent ester intermediate. Two H–bond donating residues (a conserved Trp located directly adjacent to the nucleophilic aspartate, and a second Trp or an Asn) stabilize the halide leaving group. Interestingly, many bacterial strains, which were previously not recognized as degraders of halogenated compounds, appear to possess putative haloalkane dehalogenase genes. Analysis of the genome of Mycobacterium tuberculosis H37Rv, for example, indicated the presence of three chromosomal genes encoding putative haloalkane dehalogenases, and hydrolytic dehalogenase activity was indeed detected in a number of saprophytic as well as parasitic Mycobacterium strains (Jesenska´ et al., 2000, 2002). The detection of open reading frames encoding putative dehalogenases in the genomes of two plant-symbiotic bacteria, Mesorhizobium loti MAFF303099 and Bradyrhizobium japonicum USDA110, inspired Sato et al. (2005) to analyze the activity and substrate specificity of the corresponding gene products, resulting in the identification of two haloalkane dehalogenases DmlA and DbjA with unusual substrate specificities and high catalytic activity towards b-methylated compounds. Haloalkanoic acids such as chloro- and bromoacetate, or optically pure 2-chloropropionate isomers, are used as building blocks in organic syntheses. Chloroacetate is important for the production of carboxymethyl cellulose; 2,2-dichloropropionate is the active constituent of the herbicide Dalapon. As mentioned above, haloacids have also been identified as

875

876

6

Aerobic Degradation of Halogenated Aliphatics

intermediates in the microbial degradation of halogenated alkanes. The hydrolytic dechlorination of a-halogenated low molecular weight carboxylic acids is catalyzed by haloacid dehalogenases, which can be classified into (at least) two phylogenetic groups (Hill et al., 1999). Group II enzymes selectively hydrolyze L-2-haloacids to D-2-hydroxyacids, i.e., the reaction results in the inversion of the substrate-product configuration. It may be noteworthy that the fold of group II haloacid dehalogenases, also referred to as the HAD superfamily, is completely different from the a/b-hydrolase fold of haloalkane dehalogenases, but shares a structural similarity with magnesium-dependent phosphatases and P-type ATPases (de Jong and Dijkstra, 2003). Catalysis involves nucleophilic attack by a catalytic aspartate residue, displacing the halide and resulting in a covalent enzyme-ester intermediate, and cleavage of the ester intermediate by an active-site water molecule that presumably is activated by another aspartate moiety (Li et al., 1998; Ridder et al., 1999; Schmidberger et al., 2007). In contrast, hydrolysis of 2-haloacids by group I enzymes does not involve a covalent intermediate, but proceeds via direct attack of an activated water molecule on the a-carbon of the substrate, displacing the halide (Nardi-Dei et al., 1999; Schmidberger et al., 2008). Some group I haloacid dehalogenases hydrolyze both L- and D-2-haloacids, others are specific for D-enantiomers, but the reaction also results in the inversion of the substrate-product configuration. Group I haloacid dehalogenases adopt a unique fold which till now has no structural homologues in the databases (Schmidberger et al., 2008).

4

Haloalkenes and Haloalkenoic Acids

Vinyl chloride (chloroethene) is mainly used for synthesis of its polymer, polyvinyl chloride (PVC), whereas 1,2-dichloroethene, trichlorethene, and tetrachlorethene have wide applications as industrial solvents. The chlorinated ethenes (with the exception of tetrachloroethene) can undergo fortuitous oxidation by microorganisms expressing different mono- and dioxygenase systems (reviewed in Fetzner, 1998), or productive degradation by ethene degrading bacteria. Interestingly, degradation of vinyl chloride by Nocardioides sp. JS614 is encoded on a linear plasmid. It proceeds through the ‘‘coenzyme M pathway,’’ which presumably involves a spontaneous chloride elimination step rather than a dehalogenase-catalyzed reaction (reviewed in Fetzner et al., 2007; Mattes et al., 2005) (> Fig. 4). Vinyl chloride degrading Mycobacterium spp. may use an analogous pathway (Coleman and Spain, 2003). The cis- and trans-isomers of 1,3-dichloropropene are the active ingredients of soil fumigants which have been used in agriculture to control plant-parasitic nematodes. Degradation of both the isomers is initiated by hydrolytic cleavage of the chlorine substituent at C-3 by a haloalkane dehalogenase, yielding the corresponding 3-chloroallylalcohol isomers. The latter are then oxidized to cis- or trans-3-chloracrylic acid, which are dehalogenated to malonate semialdehyde by either a cis- or a trans-specific 3-chloroacrylic acid dehalogenase (Poelarends et al., 1998) (> Fig. 5). Whereas most hydrolytic dehalogenases catalyze the cleavage of halogens bound to sp3hybridized carbon atoms, 3-chloroacrylic acid dehalogenases have the unique ability to hydrolyze – without involvement of a cofactor – the more stable vinylic carbon-halogen bond, in which the halogen is bound to an sp2-hybridized carbon atom. The cis- and trans3-chloroacrylic acid dehalogenases have a sequence identity of only about 20%, but both the proteins belong to the tautomerase superfamily. The reaction is mechanistically a hydration which – in contrast to the mechanisms of haloalkane dehalogenases and group II haloacid

Aerobic Degradation of Halogenated Aliphatics

6

. Figure 4 Hypothetical pathway of vinyl chloride degradation by Nocardioides sp. JS614 (modified from Mattes et al., 2005). (1) Vinyl chloride; (2) chloroepoxyethane; (3) 2-chloro-2-hydroxyethyl-CoM; (4) 2-ketoethyl-CoM; (5) carboxymethyl-CoM; (6) CoM-acetyl-CoA; (7) acetyl-CoA. EtnABCD, alkene monooxygenase; EtnE, epoxyalkane:coenzyme M transferase; CoA, coenzyme A; CoM, coenzyme M (2-mercaptoethanesulfonate).

. Figure 5 Degradation of trans-1,3-dichloropropene by Pseudomonas pavonaceae 170 (modified from de Jong et al., 2004). (1) 1,3-Dichloropropene; (2), trans-3-chloroallylalcohol (1-chloro-3hydroxypropene); (3) trans-3-chloroacrylic acid; (4) malonic acid semialdehyde; (5) acetaldehyde; DhaA, haloalkane dehalogenase; CaaD, trans-3-chloroacrylic acid dehalogenase.

dehalogenases – does not involve covalent catalysis (de Jong et al., 2004, 2007; Poelarends et al., 2001, 2004; Wang et al., 2003). Since the loss of halide probably is a fortuitous reaction, CaaD proteins may be considered as ‘‘accidental’’ dehalogenases (de Jong et al., 2007; Wang et al., 2003).

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Haloalcohols

2-Chloroethanol, an industrial solvent, is oxidized by Pseudomonas spp. to 2-chloracetate, which then undergoes hydrolytic dechlorination to glycolate (Strotmann et al., 1990; Stucki and Leisinger, 1983). 2,3-Dibromo-l-propanol has been used as a flame retardant, and as an intermediate in the manufacture of pesticides and pharmaceuticals. Its bacterial conversion to glycerol via epibromohydrin was already reported in 1968 by Castro and Bartnicki for a Flavobacterium sp. strain, and a series of other bacteria able to grow on short-chain haloalcohols have been isolated and identified since then. The utilization of vicinal haloalcohols depends on halohydrin dehalogenases (or halohydrin hydrogen-halide lyases), which catalyze the reversible intramolecular nucleophilic displacement of the halogen by the adjacent hydroxyl group, generating the corresponding epoxides. Halohydrin dehalogenases belong to the short-chain dehydrogenases/reductases (SDR) family (van Hylckama Vlieg et al., 2001). They occur not only in halopropanol metabolism, but also in the bacterial degradation pathway of 1,2-dibromoethane (see > Fig. 3b). Agrobacterium radiobacter strain AD1 can grow on 1,3-dichloro-2-propanol and epichlorohydrin as sole carbon source (van den Wijngaard et al., 1989) (> Fig. 6). X-ray structures of halohydrin dehalogenase HheC complexed with various ligands and kinetic studies provided insight into the catalytic mechanism and the structural basis of its high enantioselectivity for (R)-enantiomers (de Jong et al., 2003, 2005; Tang et al., 2003). In contrast to HheC, halohydrin dehalogenase from Arthrobacter sp. AD2 lacks enantioselectivity, which has been explained by a more open structure of its substrate binding pocket (de Jong et al., 2006). Halohydrin dehalogenases also catalyze the back reaction, i.e., the opening of the epoxide ring. Interestingly, HheC besides chloride, bromide, and iodide accepts other small, negatively charged nucleophiles (such as azide, nitrite, cyanide, cyanate, thiocyanate) for the epoxideopening reaction, resulting in C–N, C–C, and C–S bond formation. Since such regio- and enantioselective ring-opening of epoxides using different nucleophiles produces optically pure b-functionalized alcohols, which are versatile precursors for the synthesis of a number of

. Figure 6 Proposed pathway for the degradation of 1,3-dichloro-2-propanol in Agrobacterium radiobacter AD1 (modified from de Jong et al., 2003). (1) 1,3-Dichloro-2-propanol; (2) epichlorohydrin; (3) 1-chloro-2,3-propanediol; (4) glycidol; (5) glycerol; HheC, haloalcohol dehalogenase; EH, epoxide hydrolase.

Aerobic Degradation of Halogenated Aliphatics

. Figure 7 (Continued)

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biologically active molecules, halohydrin dehalogenases have interesting biocatalytic potential (Hasnaoui-Dijoux et al., 2008). Chloroallylalcohols are intermediates in the industrial synthesis of pesticides; cisand trans-3-chlorallylalcohol are highly nematocidal. The mineralization of 2- and 3chlorallylalcohol by Pseudomonas spp. proceeds via the corresponding 2- and 3-chloroacrylates (Belser and Castro, 1971; van der Waarde et al., 1993). Dechlorination and degradation of the latter via malonate semialdehyde may follow the pathway described for P. pavonaceae 170 (cf. > Fig. 5).

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Halogenated Alicyclic Compounds

Several insecticides, such as Lindane, Dieldrin, Chlordane, Endosulfan, Kepone, Mirex, and Toxaphene, are complex and highly chlorinated cyclic compounds. Due to their environmental persistence, toxicity, and potential for bioaccumulation along the food chain, they are banned or restricted in most countries. Most of these substances appear to be basically inert to biodegradation (Montogomery, 1997), however, Lindane (g-hexachlorocyclohexane, g-HCH) is metabolized by a number of soil microorganisms (Phillips et al., 2005). Published half-lifes of g-HCH in soil range from 4 to 6 weeks to up to 2 years, while other HCH isomers can persist for as long as 15 years. g-HCH is transformed by both aerobic and anaerobic microorganisms, but mineralization has been observed only in oxic systems (Phillips et al., 2005). The degradation pathway and its molecular basis have been studied extensively in Sphingobium japonicum strain UT26, which can utilize g-HCH as source of carbon and energy (reviewed by Nagata et al., 2007) (> Fig. 7). Three different dehalogenases LinA, LinB, and LinD mediate the key dechlorination steps. LinA, which may have evolved from a dehydratase, catalyzes two initial dehydrochlorination steps to form a diene-type intermediate (> Fig. 7). Interestingly, a- and d-HCH are also dehydrochlorinated by LinA, whereas b-HCH is not accepted as substrate. Haloalkane dehalogenase LinB, an enzyme of the a/b-hydrolase fold family, has a relatively broad substrate specificity and hence was suggested to be an attractive target for protein engineering studies to develop biocatalysts for the detoxification of different organochlorine compounds. The third dehalogenase, LinD, which is related to glutathione S-transferases, catalyzes the reductive dechlorination of the 2,5-dichlorohydroquinone intermediate (> Fig. 7). Since nearly identical lin genes have been identified in a number of HCHdegrading bacteria, it has been suggested that genes mediating HCH degradation have been spread by horizontal transfer (Nagata et al., 2007). . Figure 7 Proposed degradation pathway of Lindane (g-HCH) in Sphingobium japonicum UT26 (adapted from Nagata et al., 2007). (1) g-Hexachlorocyclohexane (g-HCH, Lindane); (2) pentachlorocyclohexene; (3) 1,3,4,6-tetrachloro-1,4-cyclohexadiene; (4) 1,2,4-trichlorobenzene; (5) 2,4,5-trichloro-2,5-cyclohexadiene-1-ol; (6) 2,5-dichlorophenol; (7) 2,5-dichloro-2,5cyclohexadiene-1,4-diol; (8) 2,5-dichlorohydroquinone; (9) chlorohydroquinone; (10) hydroquinone; (11) 3-hydroxymuconic acylchloride; (12) 3-hydroxymuconic semialdehyde; (13) maleylacetic acid; (14) 3-oxoadipic acid. GSH, glutathione. LinA, dehydrochlorinase; LinB, haloalkane dehalogenase; LinC, dehydrogenase; LinD, glutathione-dependent reductive dechlorinase; LinE, ring-cleavage dioxygenase; LinF, maleylacetate reductase.

Aerobic Degradation of Halogenated Aliphatics

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Research Needs

The biotransformation potential of microbial systems may be more extensive than our current biochemical knowledge suggests. Putative dehalogenase genes identified in genome sequences and in environmental DNA often differ significantly from related genes of bacterial strains that were obtained by classical enrichment techniques. Moreover, novel catabolic functions may be concealed in the countless genes and gene clusters that encode hypothetical proteins. Thus, the functional exploration of genome and metagenome sequences can be a promising approach to detect novel catabolic traits. Failure of microorganisms to utilize some polyhalogenated compounds is often a problem of biochemical (rather than thermodynamic) limitations, such as lack of recognition by uptake systems or regulatory proteins, or inadequate specificity and activity of catabolic enzymes. This has apparently been the case of, for example, aerobic growth on trihalogenated propanes (cf. the section on haloalkanes). Application of directed evolution techniques and other protein engineering approaches to improve key enzymes, combined with metabolic engineering of pathways, will facilitate the construction of organisms with extended or novel degradative capacities. Studies on the enantioselectivity of halohydrin and haloacid dehalogenases and on the ability of halohydrin dehalogenases to catalyze ‘‘unnatural’’ reactions suggest a large potential of these enzymes for use in biocatalysis (Janssen, 2007; Hasnaoui-Dijoux et al., 2008). Optimization of dehalogenases for biocatalytic applications and the implementation of these biocatalysts in industrial processes is another challenge for further research.

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Aerobic Degradation of Halogenated Aliphatics van den Wijngaard AJ, Janssen DB, Witholt B (1989) Degradation of epichlorohydrin and halohydrins by bacterial cultures isolated from freshwater sediment. J Gen Microbiol 135: 2199–2208. van den Wijngaard AJ, Reuvekamp PT, Janssen DB (1991) Purification and characterization of haloalcohol dehalogenase from Arthrobacter sp. strain AD2. J Bacteriol 173: 124–129. van den Wijngaard AJ, van der Kamp KWHJ, van der Ploeg J, Pries F, Kazemier B. Janssen DB (1992) Degradation of 1,2-dichloroethane by Ancylobacter aquaticus and other facultative methylotrophs. Appl Environ Microbiol 58: 976–983. van der Ploeg J, van Hall G, Janssen DB (1991) Characterization of the haloacid dehalogenase from Xanthobacter autotrophicus GJ10 and sequencing of the dhlB gene. J Bacteriol 173: 7925–7933. van der Waarde JJ, Kok R, Janssen DB (1993) Degradation of 2-chloroallylalcohol by a Pseudomonas sp. Appl Environ Microbiol 59: 528–535. van Hylckama Vlieg JET, Poelarends GJ, Mars AR, Janssen DB (2000) Detoxification of reactive intermediates during microbial metabolism of halogenated compounds. Curr Opin Microbiol 3: 257–262. van Hylckama Vlieg JET, Janssen DB (1992) Bacterial degradation of 3-chloroacrylic acid and the characterization of cis- and trans-specific dehalogenases. Biodegradation 2: 139–150.

6

van Hylckama Vlieg JET, Tang L, Lutje Spelberg JH, Smilda T, Poelarends GJ, Bosma T, van Merode AEJ, Fraaije MW, Janssen DB (2001) Halohydrin dehalogenases are structurally and mechanistically related to short-chain dehydrogenases/reductases. J Bacteriol 183: 5058–5066. Vannelli T, Messmer M, Studer A, Vuilleumier S, Leisinger T (1999) A corrinoid-dependent catabolic pathway for growth of a Methylobacterium strain with chloromethane. Proc Natl Acad Sci USA 96: 4615–4620. Verschueren KHG, Selje´e F, Rozeboom HJ, Kalk KH, Dijkstra BW (1993) Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature 363: 693–698. Wang SC, Person MD, Johnson WH Jr, Whitman CP (2003) Reactions of trans-3-chloroacrylic acid dehalogenase with acetylene substrates: consequences of and evidence for a hydration reaction. Biochemistry 42: 8762–8773. Wischnak C, Lo¨ffler FE, Li J, Urbance JW, Mu¨ller R (1998) Pseudomonas sp. strain 273, an aerobic a,odichloroalkane-degrading bacterium. Appl Environ Microbiol 64: 3507–3511. Yokota T, Omori T, Kodama T (1987) Purification and properties of haloalkane dehalogenase from Corynebacterium sp. strain m15–3. J Bacteriol 169: 4049–4954.

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Part 3

Biochemistry of Anaerobic Degradation

7 The Biochemistry of Anaerobic Methane Oxidation M. Taupp . L. Constan . S. J. Hallam Department of Microbiology and Immunology, Life Sciences Centre, University of British Columbia, Vancouver, BC, Canada [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 890 1.1 Unraveling a Methane Mystery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 890 2 2.1 2.2 2.3

Thermodynamic Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 891 Free Energy Yields . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 892 Substrate Ranges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 893 Intermediates and Electron Acceptors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 894

3 Biochemical Insights into the Anaerobic Oxidation of Methane . . . . . . . . . . . . . . . . . 896 3.1 A Conspicuous Nickel Protein . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 896 3.2 Pathway Validation and Gene Discovery . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 902 4

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 905

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_63, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: The anaerobic oxidation of methane (AOM) is a globally significant biogeochemical process that exerts a profound influence on methane flux between oceanic and atmospheric compartments of the biosphere. In marine sediments AOM occurs in a region of sulfate and methane depletion known as the sulfate–methane transition zone (SMTZ) where methane is converted to carbon dioxide and reduced products that are in turn used as electron donors in the conversion of sulfate to hydrogen sulfide and water. From a bioenergetic perspective, AOM represents a major source of maintenance energy within the SMTZ, and despite low estimated free energy yields supports a vigorous microbial metabolism. Lipid biomarker, phylogenetic stain and environmental PCR studies aimed at determining the biological component of AOM converge on microbial communities dominated by uncultivated anaerobic methaneoxidizing archaea (ANME-1, ANME-2 and ANME-3) and sulfate reducing bacteria (SRB). Specific physical associations between these groups have been observed consistent with syntrophic modes of growth. However, despite extensive mesocosm and labeling studies the precise mode of electron transfer between ANME and SRB remains unknown. Recent cultivation-independent studies of AOM communities from the Eel River Basin, Hydrate Ridge and the Black Sea have led to preliminary reconstruction of the genes and pathways mediating carbon and energy metabolism within ANME subgroups providing a genomic and proteomic basis for inferring substrate ranges, intermediates and terminal electron acceptors. The following chapter reviews biochemical aspects of AOM with special emphasis on pathway validation, electron flow and enzyme function. We consider how ANME subgroup partitioning and gene expression profiles overlap with prevailing thermodynamic models and speculate on syntrophic growth models as they relate to broader aspects of community metabolism within AOM sediments.

1

Introduction

This chapter seeks to describe recent studies focused on the substrates, intermediates, electron acceptors and enzymatic machinery underlying the anaerobic oxidation of methane (AOM) with special consideration of methanogen-related anaerobic methane oxidizing archaeal groups ANME-1 and ANME-2. For additional information on the genes predicted to encode metabolic subsystems underlying methane oxidation within ANME metagenomes the reader is referred to > Chapter 45, Vol. 3, Part 3.

1.1

Unraveling a Methane Mystery

Methane (CH4) is a potent greenhouse gas whose atmospheric concentration has increased significantly due to anthropogenic activities and fluctuated naturally over glacial and interglacial cycles. While the importance of CH4 in Earth’s climate dynamics has been well established, the biogeochemical processes regulating its oceanic cycling remain poorly understood. Despite high rates of CH4 production in marine sediments, including a number that have been targeted as petroleum reserves, net CH4 sources from the ocean to the atmosphere appear to be small. This is due in large part to the phenomenon of AOM (reviewed in (Reeburgh 2007)). In marine sediments AOM occurs in a region of sulfate (SO42
) and CH4 depletion known as the sulfate–methane interface (SMI) (Borowski et al., 1999) or sulfate–methane transition zone (SMTZ) (Iverson and Jorgensen 1985; Reeburgh 1980). Within the SMTZ, CH4 is

The Biochemistry of Anaerobic Methane Oxidation

7

converted to carbon dioxide (CO2) and reduced products that are in turn used as electron donors in the conversion of SO42
to hydrogen sulfide (H2S) and water. From a bioenergetic perspective AOM represents the major source of maintenance energy within the SMTZ and despite low estimated free energy yields supports a vigorous microbial metabolism (see Section 2.1 below). The community structure of the SMTZ is dominated by several uncultivated microbial groups, including methanogen-related anaerobic methane-oxidizing euryarchaea (ANME-1, ANME-2 and ANME-3) and sulfate-reducing bacteria (SRB), including members of the Desulfosarcinales and Desulfobulbaceae affiliated with the d subdivision of proteobacteria (reviewed in (Schleper et al., 2005; Valentine 2002)). Specific physical associations between ANME and SRB have been observed, including the formation of spheroid aggregates consisting of ANME-2 (Boetius et al., 2000) or ANME-3 (Niemann et al., 2006) surrounded by SRB. Recent fluorescent in situ hybridization (FISH) studies have revealed additional bacterial lineages associated with ANME-2, including members of the a and b proteobacteria, expanding the range of potential cooperative interactions coupled to methane oxidation (Pernthaler et al., 2008). Chemotaxonomic surveys of deep-sea sediment cores have also implicated benthic marine crenarchaea in dissimilatory methane metabolism (Biddle et al., 2006; Inagaki et al., 2006). However, direct evidence linking methane consumption to crenarchaeal growth has not been reported. Environmental genomic analysis of DNA sequences derived from AOM sediments in the Eel River Basin (ERB) (Hallam et al., 2004; Pernthaler et al., 2008), Black Sea (Kruger et al., 2003; Meyerdierks et al., 2005) and Hydrate Ridge (Meyerdierks et al., 2005) has led to the identification of numerous methanogenesis-associated genes consistent with close taxonomic and functional relationships between ANME and methanogenic archaea. Using a combination of phylogenetic anchor screening and sequence binning approaches Hallam et al., were able to reconstruct intact or nearly intact methane oxidation pathways within ANME-1 and ANME-2 metagenomes based on canonical methanogenic pathways (Hallam et al., 2004). The resulting model for AOM is consistent with a reversal of the aceticlastic pathway and the conversion of acetyl-CoA to acetate with concomitant ATP synthesis by substrate level phosphorylation. However, thermodynamic constraints associated with this mode of energy metabolism remain to be fully reconciled with syntrophic growth models. Moreover, with the exception of methyl-coenzyme M reductase purified from ANME-1 dominated bioherms in the Black Sea (Kruger et al., 2003), functional validation of this pathway in relation to electron carriers, cofactor and metabolite concentrations, protein expression or enzymatic activity remains largely unexplored.

2

Thermodynamic Considerations

When considering the bioenergetic landscape of AOM, it is instructive to look more closely at the metabolic interactions between microbes in methanogenic environments where the degradation of complex organic matter to CH4 and CO2 unfolds at the base of the redox tower (reviewed in (Zehnder and Stumm 1988)). In these settings the low free energy yields associated with the methanogenic phenotype have selected for cooperative or syntrophic modes of energy metabolism between methanogens and various primary or secondary (acetogenic) fermenting bacterial groups (reviewed in (Schink 1997; Stams et al., 2005)). For instance, primary fermentation produces molecular hydrogen that is in turn used as an

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electron source by methanogens in the reduction of CO2. Although the former reaction is endergonic under standard conditions, the efficient removal of hydrogen from the system by methanogens pushes the reaction in the oxidative direction, enabling the growth of both partners. By calculating the free energy yield across the sum of each partial-reaction, an energetically unfavorable reaction can proceed as long as syntrophic partners are capable of sharing the residual energy efficiently. This model provides a framework for developing and testing hypotheses related to ANME and SRB energy acquisition strategies.

2.1

Free Energy Yields

The estimated Gibbs Free Energy yield (DG ’) for methane oxidation coupled to sulfate reduction derived from theoretical calculations under standard conditions or using in situ profiles of substrate and product ratios ranges between
20 and
40 kJ/mol (> Table 1) (Boetius et al., 2000; Caldwell et al., 2008; Michaelis et al., 2002; Nauhaus et al., 2002; Raghoebarsing et al., 2006; Shima and Thauer 2005; Strous and Jetten 2004) þ
 CH4 þ SO2
4 þ H ¼ CO2 þ HS þ 2H2 O DG ¼
21 kJ=mol: ðThauer and Shima 2008Þ

ð1Þ

A contemporary review of AOM by Caldwell et al., suggests that recalculation of the standard equation (1) with CH4 and CO2 in the aqueous phase significantly increases the free energy yield for methane oxidation (Caldwell et al., 2008) (> Table 1). However, these calculations do not appear to factor in the effect of pH on the free energy of reaction (David Valentine, personal communication). ATP production coupled to proton translocation via the F0F1 ATP synthase requires an energetic input of between 40 and 70 kJ/mol. The c subunit composition of the rotating ring of the F0 component determines the number of protons translocated per ATP produced with a coupling number between three or four (reviewed in (Ferguson 2000)). Because the energetic cost of ATP synthesis is distributed among each proton, the minimum free energy requirement under standard conditions for a four-proton translocation system is approximately 15 kJ/mol. In cases of interspecies electron transfer the free energy yield must be distributed between syntrophic partners. Given the small free energy yield associated with methane oxidation coupled to sulfate reduction only one partner in a two part syntrophy should be capable of deriving sufficient energy from this reaction for ATP synthesis. This raises the intriguing possibility that AOM mediated by ANME and SRB is a co-metabolic activity that does not involve direct electron transfer (Schink 1997) or that structural variation within the F0 component of the archaeal or bacterial F0F1 ATP synthase supports coupling numbers greater than four. Both scenarios are consistent with the slow in situ growth rates observed for ANME populations with estimated doubling times ranging between 3 and 7 months (Girguis et al., 2003; Kruger et al., 2008b; Nauhaus et al., 2002). Contributing to the enigma of AOM energy metabolism are the rates at which methane oxidation and sulfate reduction co-occur. In (1) CH4 and SO42– are consumed in a 1:1 molar ratio. Although this ratio has been observed in a number of incubation studies (Kruger et al., 2008a; Nauhaus et al., 2002; Treude et al., 2005), sulfate reduction often exceeds methane oxidation by a factor of 2 or more (Alperin and Reeburgh 1985; Devol and Ahmed 1981; Joye et al., 2004; Orcutt et al., 2008).

The Biochemistry of Anaerobic Methane Oxidation

7

. Table 1 Gibbs Free Energy changes associated with AOM under varying environmental conditions

Reaction

DG0’ (kJ/mol DG (kJ/mol CH4) CH4)

CH4 + SO42
+ H+= CO2 + HS
+ 2H2O

Location

Reference


20

Continental shelves

Strous and Jetten (2004)a


40

Deep ocean

Strous and Jetten (2004)b

Theoretical

Shima and Thauer (2005)c

CH4 + SO42
+ H+= CO2 + HS
+
21 2H2O CH4(aq) + SO42
+ 2H+= CO2(aq) +
109 H2S + 2H2O


74

Theoretical

Caldwell et al. (2008)d

CH4 + SO42
= HCO3
+ HS
+ H2O


40

Hydrate Ridge

Boetius et al. (2000)f


30

Hydrate Ridge

Boetius et al. (2000)g


37

Hydrate Ridge

Nauhaus et al. (2002)g


20

Hydrate Ridge

Nauhaus et al. (2002)h


40

Black Sea

Michaelis et al. (2002)i

5CH4 + 8NO3
+ 8H+ = 5CO2 + 4N2 + 14 H2O


765

Theoretical

Raghoebarsing et al. (2006)j

3CH4 + 8NO2
+ 8H+ = 3CO2 + 4N2 + 10H20


928

Theoretical

Raghoebarsing et al. (2006)j

a

0.2 mM CH4 concentration 200 mM CH4 concentration c Neutral pH, CH4 and CO2 in gas phase at 0.1 MPa pressure, SO42
and HS
at 1M each d CH4 concentration of 1 M and a SO42
concentration of 2 mM e CH4 pressue 8.0 MPa, temperature 4 C, pH 7.5, 20 mM SO42
, 10 mM HCO3
and 2mM HS
f CH4 pressue 0.1 MPa, temperature 4 C, pH 7.5, 20 mM SO42
, 10 mM HCO3
and 2mM HS
g CH4 pressure: 1.1 MPa, temperature: 25 C, pH 7.5; 28 mM SO42
,30 mM HCO3
and 1 mM HS
h CH4 pressure 0.1 MPa, temperature: 25 C 10 mM SO42
, 48 mM HCO3
and 18 mM HS
i CH4 pressure 2.0 MPa j No information provided (calculated for standard conditions) b

2.2

Substrate Ranges

As described in previous sections, although ecological context for AOM is defined in large part by methane concentrations within the SMTZ, sulfate reduction rates often exceed those of methane oxidation within AOM sediments. Therefore, the possibility exists that ANME or SRB groups use alternative electron donors as part of their energy metabolism. The dissociation energy of the C–H bond in methane is + 439 kJ/mol, only the C–H bond in benzene is more stable (+473 kJ/mol). Higher alkanes including ethane, propane or butane have similar dissociation energies (ethane: +423 kJ/mol; propane: +423 kJ/mol; butane: +425 kJ/mol).

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The Biochemistry of Anaerobic Methane Oxidation

Along with methane, these compounds are found in relatively high abundance within hydrocarbon reservoirs supporting AOM and could serve as electron donors for ANME or SRB (Sassen et al., 1999; Joye et al., 2004). Radiotracer and cultivation studies indicate that fatty acids and hydrocarbon compounds including higher alkanes have the potential to fuel SRB growth within AOM sediments in the absence of methane oxidation (Joye et al., 2004; Kniemeyer et al., 2007). In contrast, Nauhaus et al., working with longer-term incubations from the Black Sea and Hydrate Ridge were unable to observe coupling between the oxidation of higher alkanes, including ethane, propane and butane, and sulfate reduction rates (Nauhaus et al., 2005). In the same study, an undefined subset of these compounds in the gas phase was inhibitory to AOM suggesting that ANME are unable to utilize higher alkanes as electron donors (Nauhaus et al., 2005). Enzyme inhibition studies in the context of energy metabolism provide a useful measure of the relative importance of specific electron donors or energy conversion pathways. Although the interpretation of inhibition experiments should be made with extreme caution due to the inherent toxicity and variable uptake of inhibitory substrates within environmental samples, used in combination with other independent growth or rate measures, they can help constrain the specific activities of target organisms or enzymatic machines within complex microbial communities. Treatment of AOM mesoscosms with inhibitory substrate analogs for methylcoenzyme M reductase (MCR), including bromoethane sulfonate has been shown to inhibit methane oxidation consistent with a role for MCR in methane activation (Alperin and Reeburgh 1985; Hoehler et al., 1994). Likewise, treatment of mesocosms with molybdate, a potent inhibitor of sulfate reduction has been shown to antagonize methane oxidation rates consistent with some form of coupling between the two processes (Nauhaus et al., 2005; Orcutt et al., 2008). However, both treatments have been observed to manifest variable effects depending on the samples under study, in some cases having little to no effect on AOM. Taken together these observations suggest that biogeographically distinct subpopulations of SRB are capable of using higher alkanes as electron donors in AOM sediments. Moreover, the coupling of methane oxidation to sulfate reduction is sensitive to inhibitors of either process alone under cell autonomous (within a single ANME cell) or syntrophic growth conditions. While it remains to be determined if specific d proteobacterial ecotypes exist solely as obligate syntrophs or in opportunistic relationships defined by substrate availability, the energy metabolism of ANME groups appears to be largely dependent on CH4 as a fuel for anaerobic growth (> Fig. 1).

2.3

Intermediates and Electron Acceptors

From the standpoint of electron flow between ANME and SRB during syntrophic growth, formate, acetate, glucose, lactate, molecular hydrogen, carbon monoxide and redox-active small molecules including phenazines and humic acids have all been proposed as potential intermediates (Nauhaus et al., 2005) (> Fig. 1). The addition of any one of these metabolites to sediment incubations does not have a significant effect on sulfate reduction rates suggesting that they do not participate in direct electron transfer between ANME and SRB (Nauhaus et al., 2005). Moran et al., have recently proposed that methylsulfides (MeSH) produced as primary end products of methane oxidation could act as electron shuttles between ANME and SRB (Moran et al., 2008) (> Fig. 1). However, support for this model remains equivocal, as the production of MeSH by ANME groups has yet to be measured in mesocosm or environmental

The Biochemistry of Anaerobic Methane Oxidation

7

. Figure 1 Schematic representation of AOM community structure and potential interactions between ANME archaea and various bacterial groups including d proteobacteria (Desulfovibrio/ Desulfococcus/Desulfobulbus), b proteobacteria (Bulkhorderia) and a proteobacteria (Sphingomonas). AOM communities consume CH4 with trace levels of methanogeneis observed (Treude et al., 2007). CO2 is both incorporated and respired (Wegener et al., 2008). Metabolites are transferred between ANME cells and surrounding community members in the form of electron shuttles, growth factors or glycocalyx. Organic acids include, acetate, formate, lactate, and amino acids, Methyl-X includes MeSH and methylamines, phenazines include any number of small redox sensitive bioactive molecules that remain undefined in AOM community settings. Terminal electron accepting processes in proposed cases of syntrophy are shown as arrows on the outer periphery of the syntrophic partner sphere. Broken lines represent theoretical couplings and question marks remain undefined.

extracts and SRB enrichment on MeSH compounds failed to support the growth of d proteobacteria associated with ANME subgroups. Extracellular electron transfer involving ‘‘nanowires’’ has also been proposed as a mechanism for interspecies electron transfer between ANME and SRB based on proximity relationships within ANME consortia (Thauer and Shima 2008). Co-metabolic processes other than direct electron transfer could help explain the apparent absence of identifiable electron shuttles between ANME and SRB. A recent study describing multi-layered microbial mats in the Black Sea suggests that the production of extracellular polymeric substances (EPS) or glycocalyx, including sugars, proteins and organic acids,

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by ANME groups could help structure AOM communities and provide a nutritional resource for SRB growth (Kruger et al., 2008a) (> Fig. 1). Related studies investigating the assimilation of methane and inorganic carbon by AOM communities indicate that autotrophic CO2 fixation by SRB is dependent on methane oxidation consistent with some form of metabolite exchange between ANME and SRB (Wegener et al., 2008). However, in this case the authors interpret the results to support a model of direct electron transfer rather than nutritional provisioning. Enrichment culture experiments suggest that AOM can be coupled to alternative electron acceptors resulting in higher energy yields, including nitrate (NO3
) or nitrite (NO2
) (Ettwig et al., 2008; Raghoebarsing et al., 2006). This process appears to be mediated by denitrifying bacteria affiliated with the NC10 candidate division in the absence of archaea (Ettwig et al., 2008) and potentially involves methane activation by a glycyl radical enzyme (Thauer and Shima 2008). Theory extends the range of suitable electron acceptors to include elemental sulfur (S0), arsenate (AsO43
), iron (III) or manganese (IV). However, in sulfate-free sediment incubations amended with NO3
, S0, iron or manganese oxides no correlation with methane oxidation was observed (Nauhaus et al., 2005) (> Fig. 1). Thauer and Shima suggest that if the canonical methanogenic pathways are reversed in AOM, the redox potential of terminal electron acceptors cannot exceed a positive threshold given the negative redox potential of the F430 prosthetic group in complex with the MCR apoenzyme (Thauer and Shima 2008). This would indeed be the case for the NO3
/NO2
couple as well as iron (III) and manganese (IV) suggesting that methane oxidation coupled to any one of these acceptors is unlikely to involve MCR in a cell autonomous manner. The S0/H2S couple however does not cross this positive threshold and therefore could formally serve as an electron acceptor in MCRmediated methane oxidation within ANME subgroups.

3

Biochemical Insights into the Anaerobic Oxidation of Methane

Neither pure cultures nor sediment-free laboratory growth conditions currently exist to facilitate the biochemical investigation of AOM. Despite these limitations progress has been made in several areas focused primarily on the properties of MCR recovered from AOM environments supporting natural enrichment of ANME biomass, including subunit identification and purification (Kruger et al., 2003), structural characterization of the F430 prosthetic group (Mayr et al., 2008) and immunocytochemical detection (Heller et al., 2008). Moreover, community proteome analysis leveraging the expanding technical resolution of Fourier transform ion cyclotron resonance mass spectrometry (FTICR-MS) detection and existing environmental genomic sequence information has begun to provide direct insight into the expressed coding potential of ANME subgroups as it relates to both central metabolic and energy conversion pathways (Constan, Page and Hallam, unpublished observations). The reader is referred to > Fig. 3 for a complete summary of recent findings as they relate to large insert genomic DNA fragments linked to ANME-1 and ANME-2 metagenomes.

3.1

A Conspicuous Nickel Protein

In methanogenic archaea, the mcr operon (mcrABCDG) encoding three structural subunits (a, b and g and the accessory genes mcrC and mcrD, gives rise to a 300 kDa heterotrimeric

The Biochemistry of Anaerobic Methane Oxidation

7

apocomplex with a2b2g2 configuration containing two binding sites that complex with two molecules of the 905 Da nickel porphyrin prosthetic group F430 to form the active holoenzyme (Ellefson and Wolfe 1981; Ermler et al., 1997). The active site region contains five posttranslationally modified amino acids within the structure of the alpha subunit (based on positioning in Methanothermobacter marburgensis): thioglycinea445, N-methylhistidinea257, S-methylcysteinea452, 5-(S)-methylargininea271 and 2-(S)-methylglutaminea400 (Ermler et al., 1997; Grabarse et al., 2000), that are with the exception of S-methylcysteine and 2-(S)methylglutamine, absolutely conserved across methanogenic lineages (Kahnt et al., 2007) (> Table 2). From a functional perspective, the MCR holoenzyme mediates the final step in methanogenesis, heterodisulfide formation between 2-thioethanesulfonate (CoM) and 7-thioheptanoyl-L-threonine phosphate (CoB) with concomitant release of CH4 (Ferry 1993). The MCR holoenzyme is a major constituent of the methanogen proteome, accounting for up 10% of total cellular protein (Bonacker et al., 1992). MCR function is integral to methanogenic energy metabolism and as such provides a robust target for the detection and classification of methanogenic archaea in the environment. In particular the mcrA subunit has been used extensively in reconstructing evolutionary relationships among methanogenic archaea, improving on the resolution of small subunit ribosomal RNA gene (SSU rDNA) trees (Lueders et al., 2001; Luton et al., 2002) (> Fig. 2). The recovery of mcr operons associated with ANME-1 and ANME-2 metagenomes (Hallam et al., 2003, 2004; Kruger et al., 2003) expanded the phylogenetic range of this marker . Table 2 Conserved and posttranslationally modified sites in methyl coenzyme M reductase subunit alphaa Posttranslationally modified amino acids 5-(S)-methyl Arg271

2-(S)-methyl Gln400

+

+

+

+

Argd

+

+

+

+ +

N-methylHisa257

Organism

thioGlya445

Methanothermobacter marburgensis

+

+

Methanococcus voltae

+

+

S-methylCysa452

Methanoculleus thermophilus

+

His

Methanocaldococcus jannaschii

+

+

c

+

Methanopyrus kandleri

+

+

c

+

+

Methanosarcina barkeri

+

+

+

+

c

ANME-1

b

d

Alad or Cysc

+ d

ANME-2

Gly

ANME-3

Glyd

His

d

Hisd

Cys

d

Cysd

c

Valc d

Glnd

Argd

Glnd

Arg

Amino acids posiitons based on Methanothermobacter marburgensis MCRA, + sign indicates modification detected (Khant et al., 2007) b No thioxylation (spontaneous hydrolysis may have occurred) c No methylation d Not determined a

897

898

7

The Biochemistry of Anaerobic Methane Oxidation

to include methanotrophic archaea (> Fig. 2) (Hallam et al., 2004). Moreover, the identification of diverse mcr alleles in the context of ANME energy metabolism posits fundamental questions regarding the accessory, structural or cofactor modifications necessary to overcome thermodynamic constraints on methane activation under variable environmental conditions. The purification of ANME encoded subunits and cofactors from bioherms in the Black Sea represent a watershed moment in this regard (Kruger et al., 2003). Working with biomass naturally enriched for the ANME-1 subgroup Kruger et al., successfully purified F430 cofactor variants with masses of 905 Da (light) and 951 Da (heavy) respectively (> Figs. 3 and > 4). While the mass spectrum of the light cofactor was identical to that of F430 from previously characterized methanogens, the mass spectrum of the heavy variant did not match that of any known methanogen (Kruger et al., 2003). Purification of MCR subunits from Black Sea samples identified two populations, one associated with the heavy variant termed Ni-protein I and the other associated with the light cofactor termed Niprotein II (Kruger et al., 2003). The relative proportions of both proteins was very high, with Ni-protein I representing approximately 7% (Kruger et al., 2003) and Ni-protein II representing approximately 3% (Mayr et al., 2008) of total protein extracted from environmental samples. Mass spectrometric analysis of peptide fragments associated with Ni-protein I identified them as ANME-1 encoded based on comparison to environmental mcr genes recovered from ANME metagenomes (Hallam et al., 2003; Kruger et al., 2003). Moreover, the identification of F430 cofactor variants across different AOM habitats including Hydrate Ridge and the Baltic Sea suggests that they have potential utility as biomarkers for the detection of methanotrophic archaea when biomass is not limiting (> Figs. 3 and > 4). If it is determined that ANME populations enriched in specific mcr alleles are associated with an abundance of the lighter F430 cofactor, the relative proportion of heavy and light cofactor variants could be used as a proxy for ANME subgroup partitioning in the absence of bona fide methanogens. The solution structure of the heavy and light F430 cofactor variants from the Black Sea were recently solved by NMR spectroscopy (Mayr et al., 2008). While the light cofactor was found to be identical to F430 from methanogens, the heavy cofactor was found to contain a methylthio-group substituent at position C172 of the F430 pentamethyl ester (Mayr et al., 2008). The physicochemical properties of this modification in relation to MCR catalysis remain to be investigated. Although ANME encoded MCR subunits are homologous to cognate subunits encoded in the genomes of methanogenic archaea, a closer examination of the predicted amino acid sequences of ANME-1 encoded MCRA subunits (group a/b in > Fig. 2) reveals several important differences in and around the active site of the enzyme with the potential to alter active site geometry in accordance with the structure of the modified F430 (> Table 2). These include a substitution of glutamine (Gln) to valine (Val) at the corresponding a400 position in the ANME-1 MCRA subunit and the insertion of a cysteine-rich motif (CCXXXXCXXXXXC) spanning positions a403–a415 immediately adjacent to the active site (Hallam et al., 2003; Kruger et al., 2003). Mayr et al., hypothesize that the Gln > Val substitution in the ANME-1 MCRA subunit alters active site geometry in order to accommodate the modified F430 (Mayr et al., 2008). They further speculate that the cysteinerich motif, although dissimilar to known iron–sulfur clusters, could play a role in proton or electron transfer reactions from the surface of the protein to the active site in order to modulate the Ni-oxidation state of the F430 prosthetic group (Mayr et al., 2008). These sequence features appear to be specific to ANME-1 encoded mcr alleles, suggesting that they along with the modified F430 are not absolutely required for methane activation in the context of AOM.

. Figure 2 (Continued)

The Biochemistry of Anaerobic Methane Oxidation

7

899

900

7

The Biochemistry of Anaerobic Methane Oxidation

Until very recently, the biochemical investigation of ANME MCR cofactor and subunit composition has been based on strong but circumstantial linkages between biomass extracts and metagenomic sequence information recovered from AOM environments naturally enriched for ANME-1 subgroups. Quantification of gene expression of specific mcr alleles within ANME subgroups remains to be determined at the cellular level using transcriptomic or proteomic approaches. However, recent immunocytochemical detection studies using the immunogold labeling technique have for the first time detected MCR expression within ANME-1 and ANME-2 cells recovered from bacterial mats in the Black Sea (Heller et al., 2008) (> Figs. 3 and > 4). Antibodies raised against MCR from Methanococcus vannielii cross reacting with total protein extracts from the mats were used to probe thin sections of resin embedded cells recovered from different regions of the mat enriched for either ANME-1 or ANME-2 subgroups (Heller et al., 2008). The resulting localization and enumeration of immunogold particles was consistent with high levels of MCR expression throughout the mat structures, comparable to log phase growth expression in several methanogenic lineages (Heller et al., 2008). However, the level of resolution within ANME cells was not sufficient to localize the enzyme complex to discrete subcellular compartments or cytosolic structures.

. Figure 2 A phylogenetic comparison of genes encoding methyl coenzyme M reductase subunit alpha (mcrA) and small subunit ribosomal RNA (SSU rDNA) derived from environmental clones and primary methanogenic lineages. For purposes of tree construction and presentation, an HKY evolutionary model was used in maximum likelihood analysis and bootstrap values based on 100 replicates each are shown for branches with greater than 50% support. Methanopyrus kandleri was used as the out group reference. Scale bars represent 0.1 nucleotide substitutions per site. Accession numbers for corresponding mcrA sequences are as follows: Methanosarcina acetivorans (AE010299.1), Methanosarcina barkeri (CP000099.1), Methanococcoides burtoni (CP000300.1), Methanothermus fervidus (J03375.1), Methanospirillum hungatei (CP000254.1), Methanococcus jannaschii (L77117.1), Methanopyrus kandleri (AE009439.1), Methanocorpusculum labreanum (CP000559.1), Methanoculleus marisnigri (CP000562.1), Methanococcus maripaludis (BX950229.1), Methanosarcina mazei (AE008384.1), Methanosphaera stadtmanae (CP000102.1), Methanosaeta thermophila (CP000477.1), Methanococcus vannielii (CP000742.1), Methanococcus voltae (X07793.1), Methanothermus thermoautotrophicus (U10036.1), ANME-1 subgroup a/b representative (AY324369.1), ANME-2 subgroup c/d representative (AB362199.1), AOM-affiliated subgroup e representative (DQ521858.1), and ANME-3 subgroup f representative (AM407730.1). Accession numbers for corresponding SSU rDNA sequences are as follows: Methanosarcina acetivorans (M59137.2), Methanosarcina barkeri (NC_007355), Methanococcoides burtoni (NC_007955), Methanothermus fervidus (M59145.1), Methanospirillum hungatei (NC_007796), Methanococcus jannaschii (M59126.1), Methanopyrus kandleri (AB301476.1), Methanocorpusculum labreanum (NC_008942), Methanoculleus marisnigri (NC_009051), Methanococcus maripaludis (U38941.1), Methanosarcina mazei (AE008384), Methanosphaera stadtmanae (NC_007681), Methanosaeta thermophila (NC_008553), Methanococcus vannielii (NC_009634), Methanococcus voltae (NZ_ABHB01000001), Methanothermus thermoautotrophicus (AY196661.1), ANME-1a representative (AY053468.1), ANME-1b representative (AJ578089.1), ANME-2a representative (AF354130.1), ANME-2c representative (AY323221.1), and ANME-3 representative (AJ578119.1).

cdh

acs

N5-methyltetrahydromethanopterin coenyzme M methyltransferase

methyl coenzyme M reductase

heterodisulfide reductase

carbon monoxide dehydrogenase/aceryl-CoA synthase

ADP-forming acetyl-CoA synthetase

4

5

6

7

8

. Figure 3 (Continued)

hdr

F420-dependent methylenetetrahydromethanopterin dehydrogenase

F420-dependent N5, N10-methylenetetrahydromethanopterin reductase mer

mtr

N5, N10-methenyltetrahydromethanopterin cyclohydrolase

3

Activity measured

mcr

mtd

mch

ftr

formylmethanofuran-tetrahydromethanopterin formyltransferase

fmd

2

Gene(s) identified

Black sea

Protein identified

Protein purified

locus ANME-1 ANME-2c ANME-1 ANME-2

formylmethanofuran dehydrogenase

Enzyme name

1

Step

Eel river

Cofactor identifed

ANME-1 ANME-2a ANME-2c ND

Protein localized

ND

Hydrate ridge

AOM community

ANME-1

ND

Baltic sea

The Biochemistry of Anaerobic Methane Oxidation

7 901

902

7 3.2

The Biochemistry of Anaerobic Methane Oxidation

Pathway Validation and Gene Discovery

The identification of expressed mcr alleles and F430 cofactor variants associated with ANME groups provides a functional genomic anchor point for the reconstruction of methane oxidation pathways. In methanogenic archaea utilizing the hydrogenotrophic pathway, six cofactors and more than 31 genes encoding eight enzymatic steps are needed to reduce CO2 with H2 culminating in methane release and energy production (a more in-depth examination of the component parts can be found in > Chapter 45, Vol. 3, Part 3). Beyond the core methanogenic pathway, the genetic information needed for the biosynthesis of cofactors, respiratory chains and in the regulation of methanogenic gene expression (transcription, translation and posttranslational modification) further expands the genomic repertoire necessary to manifest this phenotype. Therefore, each component part of each methanogenic subsystem represents a functional reference for identifying conserved pathway architecture between methanotrophic and methanogenic archaea. For instance, in the Black Sea study that purified the heavy and light ANME MCR variants, the activities of several methanogenic enzymes including FTR, MCH and MTD (> Figs. 3 and > 4 steps 2–4) was also detected in batch preparations consistent with the expression of these gene products by undefined ANME groups (Supplementary on-line material (Kruger et al., 2003)). In addition to isolation or activitybased measurements, direct comparison of ANME metagenomes to the genomes of cultivated methanogenic reference strains has the potential to help explain functional variation based on the absence or under representation of canonical methanogenesis genes among and between ANME subgroups or environmental samples (Hallam et al., 2004; Pernthaler et al., 2008). In the absence of gene expression or functional data, metagenomic findings should be interpreted with caution. Therefore, given the dearth of cultivated reference strains for AOM, community or strain resolved expression studies are necessary to move beyond the coding potential provided by metagenomic surveys alone. Transcriptomic or proteomic approaches enable the validation of gene models and pathways predicted during the annotation phase of metagenome analysis, and both have the capacity to identify expressed hypothetical genes with novel or conserved functions. Recent progress has been made in this regard using liquid chromatography (LC) FTICR-MS on batch protein preparations from AOM sediments. Shotgun proteomics reveals ANME encoded peptides covering enzymatic components for seven out of eight steps mediating hydrogenotrophic methanogenesis and five out of five steps mediating aceticlastic methanogenesis, consistent with a reversal of one or both pathways in ANME energy metabolism (Constan, Page and Hallam, unpublished observations) (> Figs. 3, > 4 and > 5). Moreover, functional profiling of the ERB community proteome using categories defined by clusters of orthologous groups (COG) reveals an abundance of

. Figure 3 A functional overview of canonical methanogenic pathway components identified within methane oxidizing communities with special emphasis on ANME-1 and ANME-2 subgroups. For purposes of gene identification, only loci linked to large insert genomic clones binned to ANME groups are included (Hallam et al., 2004). The term ND designates undetermined taxonomic affiliation. References cited include ERB (Hallam et al., 2004), Black Sea (Heller et al., 2008; Kruger et al., 2003; Mayr et al., 2008; Meyerdierks et al., 2005), Hydrate Ridge (Kruger et al., 2003; Meyerdierks et al., 2005) and Baltic Sea (Kruger et al., 2003). Structural characterization of the ANME-1 F430 cofactor variant is included in the cofactor identification label.

7

The Biochemistry of Anaerobic Methane Oxidation step

CO2 1

2e–

fmd

2e–

formyl-MF 2

ftr

2e–

formyl-H4MPT 3

mch

[CO] methenyl-H4MPT

4

2e–

mtd

methylene-H4MPT

CH2O H4MPT

5

6

Methylamines

2e–

2e–

2e–

methyl-H4MPT HS-CoM mtr ?

Methanol 7

Methylsulfides

methyl-S-CoM HS-CoB 2e– 2e– CH4 CH4 CoM-S-S-CoB e–– e

8

acetyl-CoA acs

cdh

ATP ATP Acetate mcr

hdr

HS-CoM + HS-CoB Gene(s) identified

Activity measured

Protein identified

Protein purified

Protein localized

Cofactor identifed

. Figure 4 A conceptual map of ‘‘reverse’’ methanogenesis pathways based on canonical methanogenesis genes in relation to cultivation-independent biochemical and gene expression data. The mer gene, mediating step 5 in the hydrogenotrophic pathway has not yet been definitively linked to ANME genotypes and is therefore depicted in gray. Genes (black), cofactors (purple), protein products (orange or purple) or activities (red) identified in one or more AOM communities including Eel River Basin, Hydrate ridge, Black Sea and Baltic Sea are depicted in the form of nested circles. Refer back to > Fig. 2 as a guide to enzyme names locus abbreviations.

energy conversion (C), cofactor (H) and amino acid biosynthetic (E) pathway components in addition to active transposases within the replication, recombination and repair category (L) (> Fig. 5). The overrepresentation of C, H and E categories is consistent with the hypothesis that ANME groups subsist close to the thermodynamic limits of life, investing most of their maintenance energy in the synthesis of the enzymatic machinery necessary to build cofactors and carry out C1-transfer reactions. The presence of active transposases suggests that lateral gene transfer between community members could play an important role in niche adaptation and ecotype partitioning among and between ANME and SRB groups.

903

7

The Biochemistry of Anaerobic Methane Oxidation

. Figure 5 (Continued)

904

The Biochemistry of Anaerobic Methane Oxidation

4

7

Research Needs

Although syntrophy remains the dominant paradigm for interpreting the energy metabolism of AOM, after decades of mesocosm and labeling studies, the metabolites mediating interspecies electron transfer within ANME consortia remain unknown. This is due in part to a general lack of knowledge regarding the chemical ecology of AOM, including in situ concentrations of postulated reaction intermediates and further confounded by the lack of pure cultures or cocultures for in-depth physiological or biochemical investigations. Future stable isotopic labeling and incubation studies should address this deficit by focusing on the metabolomic signatures of AOM across ecological gradients in order to resolve inconsistencies between existing thermodynamic models and in situ modes of energy conversion. While cultivationindependent biochemical investigations have begun to reveal the structural and functional properties of a subset of methanogenic loci within ANME dominated ecosystems, the underlying mechanisms of ‘‘reverse’’ methanogenesis with respect to enzymatic form and function are confounded by inherent biomass limitations and allelic heterogeneity associated with the majority of AOM communities. Advances in gene expression profiling, including transcriptomic and proteomic analysis at both community and strain resolved levels promises to validate and expand upon existing pathway models for AOM and provide strain resolved functional targets for downstream gene expression studies in appropriate heterologous host systems. In particular, the development of genetic tools for high-throughput or directional screening of putative methanogenic loci recovered on large insert genomic fragments using genetically tractable methanogens has great potential to enable functional and structural studies of methane oxidation pathway components.

Acknowledgments We would like to thank the Canadian Foundation for Innovation, the British Columbia Knowledge Development Fund and the National Sciences and Engineering Research Council (NSERC) of Canada for supporting ongoing studies on the anaerobic oxidation of methane. L.C. was supported by a fellowship from NSERC and M.T. was supported by fellowships from Deutsche Forschungsgemeinschaft (DFG) Germany and the TULA foundation funded Centre

. Figure 5 A workflow and data summary for ANME and SRB community proteomes based on shotgun proteomic analyses of ERB sediments. (a) A workflow for protein identification from environmental samples. Whole proteins are extracted directly from sediments, tryptically digested and submitted to LC-MS-MS (LTQ-FT Thermo). Peptide assignments are made using SEQUEST against a clustered metagenomic database to remove sequence redundancy (Norbeck and Hallam, unpublished observations). Peptide identifications are made on the basis of the Washburn and Yates criteria (Washburn et al., 2001) and hold a peptide prophet score 0.8 (Keller et al., 2002). (b) A bubble chart gene content summary of functional COG categories represented in the Eel River whole genome shotgun (wgs) library in comparison to observed peptides and proteins. Scale bar represents percent total within each COG category at each level of analysis (53,212 wgs genes with bit score cut-off >80), peptides (2,500) or proteins (527 with at least two peptide assignments).

905

906

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The Biochemistry of Anaerobic Methane Oxidation

for Microbial Diversity and Evolution. We would also like to thank Heather Mottaz, Angela Norbeck and Ljiljana Pasa-Tolic at the US Department of Energy (DOE) funded Environmental Molecular Sciences Laboratory (EMSL) located at Pacific Northwest National Laboratory (PNNL) for proteomics and bioinformatics capacity and our most excellent friends and colleagues David Walsh, Antoine Page and Leonard Foster for fruitful discussions and advice.

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8 Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes F. Widdel . O. Grundmann Max Planck Institute for Marine Microbiology, Bremen, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 910

2

Uptake of Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 911

3 Degradation of Alkanes via Alkylsuccinates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 913 3.1 Activation Yielding Alkylsuccinates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 913 3.2 Degradation of Alkylsuccinates to Acetyl-CoA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 918 4

Other Degradation Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 919

5

Terminal Oxidation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 919

6

Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 921

7

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 921

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_64, # Springer-Verlag Berlin Heidelberg, 2010

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Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

Abstract: Alkanes (saturated hydrocarbons) are naturally wide-spread compounds that are chemically unreactive. Whereas aerobic alkane-utilizing microorganism have been investigated since the early 20th century, anaerobic alkane-degraders became known relatively recently. Several nitrate- or sulfate-reducing bacteria able to growth with alkanes as sole organic substrates have been described. Furthermore, anaerobic alkane degradation was demonstrated in enriched bacterial communities under conditions of nitrate reduction, sulfate reduction, or methanogenesis. This article presents an overview of the anaerobic metabolism of non-methane alkanes. The anaerobic activation of alkanes in the absence of oxygen presents a ‘‘metabolic challenge’’. Metabolite analyses suggest that many anaerobic alkane degraders make use of a radical-catalyzed carbon-carbon addition to fumarate yielding alkylsuccinates; usually, the subterminal carbon atom of the alkane is activated, resulting in (1-methylalkyl)succinates. Isotope labeling studies suggest a further metabolism of (1-methylalkyl)succinylCoA via carbon skeleton rearrangement and decarboxylation yielding 4-methyl-branched fatty acid thioesters; the latter can undergo b-oxidation. The pathway involves a delicate stereochemistry. Furthermore, there are hints at an alternative, still unexplored pathway for anaerobic alkane degradation with the introduction of a carbon dioxide-derived carboxyl group at carbon-3.

1

Introduction

Alkanes or saturated hydrocarbons are wide-spread organic compounds in the environment which originate from natural gas and oil seeps (e.g. in geothermally active marine sediments), from the anthropogenic use of petroleum and its refined products, and as metabolites from living organisms. The most abundant alkanes are those with straight chains (C-even and C-odd n-alkanes, formula: CnH2n+2). Petroleum contains in addition various branched alkanes as well as cyclic alkanes (cycloalkanes; formula of monocyclic alkanes, CnH2n). Apart from the microbial formation of methane, the geochemical and biological reactions leading to alkanes are not fully understood from a mechanistic point of view (chapters by Petsch, Formolo, McInnerney et al., and Ferry, this volume). Alkanes, belong to the chemically least reactive organic compounds. This was formerly expressed by the designation ‘‘paraffins’’ (from Latin ‘‘parum affinis", expressing ‘‘not very related’’ or ‘‘no affinity’’). Nevertheless, numerous microorganisms can grow with alkanes as sole organic substrates. Channeling of alkanes into the metabolism requires the introduction of a functional group. The crucial step in functionalization of a saturated hydrocarbon (viz. that contains only sp3-carbon atoms) is the initial cleavage of a strong, apolar C
H-bond. Aerobic alkane-degrading microorganisms, which are known since the early 20th century, make use of a highly reactive transition metalcoordinated oxygen atom derived from O2 by partial reduction (monooxygenase reaction; chapters by Rojo, Schubert, and Widdel and Musat, this volume). Such an O-atom inserts into a C
H-bond via its ‘‘disruption’’ to yield a hydroxyl group (alcohol) in a highly exothermic and exergonic reaction. The chemical inertness of alkanes together with the fact that aerobic alkane-degrading microorganisms always make use of a highly reactive O2-derived oxygen species has for a long time favored the view that alkanes are biologically inert in the absence of oxygen. During the 1990s and thereafter, however, an increasing number of pure cultures was shown to definitely grow with alkanes as the only organic substrates that were oxidized with nitrate (Ehrenreich et al., 2000) or sulfate (Aeckersberg et al., 1991, 1998; Rueter et al., 1994; So and Young 1999; Cravo-Laureau et al., 2004; Davidova and Suflita 2005).

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

8

The isolated anaerobic alkane-degrading microorganisms were not closely related to established species and represented novel species or genera. Early reports of an anaerobic alkane degradation by established bacterial species such as Desulfovibrio or Pseudomonas strains could not be repeated in later studies (for overview see Aeckersberg et al., 1991, 1998; Ehrenreich et al., 2000). Also enriched microbial communities were shown to degrade alkanes anaerobically with sulfate (Kropp et al., 2000; Gieg and Suflita 2002), nitrate (Callaghan et al., 2009), or under conditions of methanogenesis (Zengler et al., 1999; Anderson and Lovley 2000; Jones et al., 2008). All n-alkanes so far shown to be degraded by anaerobic microorganisms were completely oxidized to carbon dioxide, or completely converted to carbon dioxide and methane, according to the following general equations: þ 5 Cn H2nþ2 þ ð6n þ 2Þ NO
3 þ ð6n þ 2ÞH ! 5n CO2 þ ð3n þ 1Þ N2 þ ð8n þ 6Þ H2 O

ð1Þ

þ 4 Cn H2nþ2 þ ð3n þ 1Þ SO2
4 þ ð6n þ 2ÞH ! 4n CO2 þ ð3n þ 1Þ H2 S þ ð4n þ 4Þ H2 O ð2Þ

4 Cn H2nþ2 þ ð2n
2Þ H2 O ! ðn
1Þ CO2 þ ð3n þ 1Þ CH4

ð3Þ

Enriched populations were also shown to degrade the branched-chain alkane pristane with nitrate (Bregnard et al., 1997), and cyclic, mainly alkyl-substituted alkanes with sulfate (Townsend et al., 2004) as electron acceptors. The anaerobic metabolism of alkanes so far has been investigated to a lesser extent than the anaerobic metabolism of the alkylbenzenes, toluene and ethylbenzene (chapters by Boll and Heider, and Tierney and Young, this volume). The reason is presumably that anaerobic growth with n-alkanes is even slower than that with the alkylbenzenes. Also, application of alkanes as very hydrophobic compounds that are less water-soluble than toluene or ethylbenzene often prevents cultivation of cells homogeneously in the medium; cells rather grow attached to the alkane phase. Formulated anaerobic pathways and reactions of alkanes are largely based on the chemical identification of metabolites, substrate isotope labeling studies, and analogies (on the level of metabolites or identified genes) to steps in the anaerobic degradation of toluene, and to long-known catabolic pathways of aliphatic acids (e.g. b-oxidation). The initial anaerobic reaction of the alkane, the activating step, has been of particular interest. Because a use of reactive oxygen species for alkane hydroxylation is excluded or unlikely under anoxic conditions, the anaerobic mechanisms for alkane functionalization are expected to represent unusual, unprecedented reactions of chemically non-activated carbon atoms (C-atoms remote from hetero-atoms or p-electron systems). The present chapter presents an overview of the metabolism of non-methane alkanes. The anaerobic oxidation of methane, which is performed by special archaeal-bacteria consortia and involves biochemically different mechanisms (related to those in methanogenesis) not encountered in the anaerobic degradation of other hydrocarbons, is treated in separate chapters (chapters by Hallam, Boll and Heider, and Widdel and Musat, this volume).

2

Uptake of Alkanes

Gaseous alkanes (C1 to C4) have noticeable solubility in water (> Fig. 1) and like other gases (H2, N2, O2) are assumed to diffuse freely into the cell. Liquid and solid alkanes as very hydrophobic substances are poorly soluble to essentially insoluble in water. They form

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Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

. Figure 1 Solubility of alkanes and some aromatic hydrocarbons (for comparison) in water. Drawn according to Wilhelm et al., (1977) and Eastcott et al., (1988).

buoyant layers or droplets, and the dissolved concentrations that are in equilibrium with the alkane phases (the maximum or saturation concentrations) dramatically decrease with increasing chain length (> Fig. 1). Hence, supply of the alkane to a planktonic (free living) cell in the aqueous medium is usually strongly limited by diffusive substrate transfer. Access of cells to alkanes (and other insoluble hydrocarbons) can in principle be improved (i) by adherence to the alkane phase enabled by hydrophobic cell surface structures or (ii) by production of amphiphilic emulsifying compounds (biosurfactants) that form micelles including alkane molecules so as to increase their aqueous ‘‘concentration’’ and diffusion to cells. Whereas both mechanisms are known in aerobic alkane degraders, cells of anaerobic alkane degraders so far were only observed to make use of an attachment to the alkane phase for better access (example of a hexadecane-utilizing sulfate reducer shown in > Fig. 2). Reported exceptions are sulfate-reducing bacteria growing with propane and butane (Kniemeyer 2007), and the denitrifying Betaproteobacterium, strain HxN1, growing with n-hexane. Growth of the latter in a planktonic mode, without attachment to the alkane phase, is explained by a certain solubility of n-hexane (> Fig. 1). There is generally no evidence for an active uptake of alkanes. Alkanes are assumed to partition in the cytoplasmic membrane and to diffuse from there to the active sites of the activating enzymes that may be also membrane-associated.

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

8

. Figure 2 Cells of the sulfate-reducing strain Hxd3, tentatively named Desulfococcus (formerly Desulfobacterium) oleovorans adhering to a droplet of n-hexadecane (Aeckersberg et al., 1991). The direct contact obviously enables passive uptake of the insoluble hydrocarbon via diffusion.

3

Degradation of Alkanes via Alkylsuccinates

3.1

Activation Yielding Alkylsuccinates

In a sulfate-reducing enrichment culture grown with n-dodecane (Kropp et al., 2000) and a denitrifying strain anaerobically grown with n-hexane (Rabus et al., 2001), alkylsuccinates with alkyl substituents matching the alkane substrate were identified. This and the obvious analogy to the formation of benzylsuccinate during anaerobic degradation of toluene (Biegert et al., 1996; Beller and Spormann 1997) suggested an alkane activation by carbon-carbon addition to fumarate as co-substrate. The usual site of attack is apparently the subterminal carbon (C-2) atom of the alkanes (> Fig. 3a). However, additionally identified metabolites indicated that n-hexane may be also activated at the C-3 atom (Rabus et al., 2001) and propane at the C-1 atom (Kniemeyer et al., 2007), most likely in side-reactions. The EPR spectroscopic measurement of a radical specifically in alkane-grown cells (Rabus et al., 2001), like in toluene-grown cells of other denitrifiers (Krieger et al., 2001; Verfu¨rth et al., 2004), was strongly supportive of a reaction catalyzed by a radical mechanism. Homolytic (radicalic) C
H-bond cleavage is indeed the most conceivable mechanism for alkane activation, as in aerobic degradation (> Chapter 1, this volume). Despite the significant energetic ‘‘barrier’’ of C
H-bond cleavage (absolute value, 400 kJ mol
1), the total reaction is exergonic (Rabus et al., 2001): H3 C
alkyl þ Fumarate ! ð1-MethylalkylÞsuccinate; DG 0 ¼
33 mol
1

ð4Þ

913

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Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

. Figure 3 (Continued)

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

8

. Figure 3 Proposed reactions in the anaerobic degradation of n-alkanes in most anaerobes studied so far. (a) Overall pathway. The initial reaction with fumarate (1; details with radical not shown) involves abstraction of an H-atom (H▪) yielding a secondary alkyl radical (R
CH2
▪CH
CH3) that adds to fumarate. The (1-methylalkyl)succinyl radical is then ‘‘saturated’’ with the initially abstracted H-atom to yield the first stable intermediate, (1-methylalkyl)succinate. The stereogenic centers are indicated by asterisks. A stereogenic center in the case of deuterium labeling of fumarate is also indicated (x). One deuterium exchanges with hydrogen from water by an unknown mechanism. After an assumed activation to the thioester with free CoA or via a CoA-transferase (2), the C-skeleton is rearranged (3) to allow decarboxylation or transcarboxlyation (4). Dehydrogenastion of (1-methylalkyl)succinyl-CoA (5) specifically removes the remaining deuterium. (b) Potential stereoisomers formed during reaction of n-hexane with fumarate. The possible positions of the C1-carbon around the C2-C3 axis of the n-hex-2-yl radical is indicated by dotted lines.

The assumption of a radical-catalyzed alkane addition to fumarate, analogous to toluene activation, was further corroborated by the analysis of genes and relationships of deduced proteins in anaerobic alkane degraders (Callaghan et al., 2008; Grundmann et al., 2008). Proteins formed specifically during growth on alkanes were shown to exhibit sequence similarity to the large subunit of benzylsuccinate synthase (today mostly termed BssA), the anaerobic toluene-activating enzyme (Coschigano et al., 1998; Leuthner et al., 1998; Shinoda et al., 2005). The alkane-specific proteins as well as benzylsuccinate synthase contain an RVXG-motif (close to the C-terminus) that is characteristic of so-called glycyl radical enzymes. Groundbreaking insights into these unique enzymes had been obtained from the study of pyruvate-formate lyase (Knappe et al., 1984). In this, the radical was shown to reside on the glycine [
HN
▪CH
CO
] in the RVXG-motif of the enzyme in its active state (Wagner et al., 1992; Becker et al., 1999). The glycyl radical is obviously a storage radical

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8

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

which converts a cysteine in the active site to a cysteyl (cysteine thiyl) radical [
HN
(▪S
CH2
)CH
CO
]; the latter then abstracts a hydrogen atom from the substrate to initiate its reaction. The product radical finally accepts the hydrogen atom from the cysteine that regains its radical state for the next catalytic round. The apparent glycyl radical proteins from the anaerobic alkane degraders are assumed to represent the large subunits of the alkaneactivating enzyme termed alkylsuccinyl synthase (Ass; Callaghan et al., 2008) or (1-methylalkyl)succinate synthase (Mas; Grundmann et al., 2008). Two smaller proteins with some similarity to those found in native benzylsuccinate synthase (BssABC) may be part of the native Ass or Mas enzymes. But additional subunits in the alkane-activating enzymes cannot be excluded on the basis of the present studies. An enzyme closely related to Ass or Mas, and also to Bss, is the assumed enzyme that activates 2-methylnaphthalene by methyl group addition to fumarate (Nms; Musat et al., 2009). Somewhat more distantly related glycyl radical enzymes are, like pyruvate-formate lyase, 2-oxobutyrate-formate lyase (Sawers et al., 1998), anaerobic (also: class III) ribonucleotide reductase (Jordan and Reichard, 1998), the B12-independent glycerol dehydratase (forming 3-hydroxypropionaldehyde; O’Brien et al., 2004), and 4-hydroxylphenylacetate decarboxylase (forming p-cresol; Selmer and Andrei, 2001). Apart from the glycyl/cysteyl radical mechanism, the reactions catalyzed by these enzymes differ significantly from hydrocarbon addition to fumarate as a co-substrate. > Fig. 4 shows sequence relationships of several glycyl radical enzymes as well as the arrangement of the genes that encode the assumed alkane-activating enzymes. The glycyl radical is introduced by a protein-activating enzyme which generates the radical by a reductive one-electron cleavage of S-adenosylmethionine (SAM) leading to a 50 -deoxyadenosyl radical (Buckel and Golding 2006; Frey et al., 2008) and methionine. Glycyl radical enzymes may therefore be termed secondary SAM-dependent radical enzymes (Buckel and Golding 2006), because the generated radical once introduced into the polypeptide chain allows an indefinite number of catalytic cycles without further involvement of SAM. In primary SAM-dependent radical enzymes, the 50 -deoxyadenosyl radical formed from SAM reacts directly with the compound to be metabolized (Buckel and Golding 2006; Frey et al., 2008). As for benzylsuccinate synthases (Coschigano et al., 1998; Leuthner et al., 1998), there are also potential (predicted) protein-activating enzymes for the glycyl radical enzymes for alkane activation. The protein-activating enzymes contain a CxxxCxxC motif without a fourth cysteine; this has been observed in all SAM-enzymes investigated so far (Frey et al., 2008). The three cysteines bind three iron ions of a special Fe4S4-cluster, leaving one ‘‘free’’ iron to react with SAM. The activation reaction of alkanes involves some peculiarities. The energy for homolytic C
H-bond dissociation at a secondary carbon (absolute value, 400 kJ mol
1) or primary carbon (414 kJ mol
1) of an alkane is higher than at the methyl group of toluene (368 kJ mol
1). The products, (1-methylalkyl)succinates, harbor two chiral carbon atoms, in comparison to one chiral carbon atom in benzylsuccinate. Whereas specific formation of the enantiomer, R(+)-benzylsuccinate (Beller and Spormann 1998; Leutwein and Heider 1999) as well as transfer of the initially abstracted H-atom to the same face of fumarate (Qiao and Marsh 2005) has been demonstrated for toluene activation, the stereochemistry in the case of alkane activation is far less clear. The formation of diastereomers of methylalkylsuccinates (yielding separate elution peaks on non-chiral gas chromatographic columns) indicated relaxed stereo-specificity at least at one of the carbon atoms forming the new C
C-bond (C-2 of the alkane, or methenyl carbon of fumarate). All possibilities for stereoisomer formation are indicated in > Fig. 3b. Another finding not explained so far was the complete

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

8

. Figure 4 Relationships on the molecular level of the assumed enzyme for anaerobic activation of n-alkanes to other glycyl radical enzymes. (a) Gene-derived amino acid tree of the catalytic subunits carrying the glycyl radical. (b) Genes potentially involved in the anaerobic activation of n-alkanes in the denitrifying Betaproteobacterium, strain HxN1, and the sulfate-reducing bacterium, strain AK-01 (containing two gene clusters), in comparison with the genes for toluene activation (an alternative designation of bss is tut; see Coschigano et al., 1998). Further genes in the operon with non-predicted function are not included. Genes encoding the catalytic subunit carrying the glycyl radical are indicated in black, the genes for the radical-generating enzyme (protein-activating enzyme) in grey. The genes termed here b1 and b2 have not been designated in the original publication, but they are apparent orthologs of masE (and bssB) which is assumed to encode the b-subunit (second largest subunit).

917

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Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

replacement of the D-atom by an H-atom at the tertiary carbon atom of the succinate moiety of (1-methylpentyl)succinate if deuterated fumarate (d2-fumarate) was supplied in addition to n-hexane (Rabus et al., 2001); the (1-methylpentyl)succinate carried only one of the initially two D-atoms (> Fig. 3a). There is no evidence so far for such a ‘‘labile’’ hydrogen atom in the apparently analogous formation of benzylsuccinate. Each anaerobic alkane degrader investigated so far exhibited a relatively narrow substrate range with respect to alkanes. For instance, the sulfate-reducing strain BuS5 can only grow with propane and n-butane (Kniemeyer et al., 2007), and the denitrifying strain HxN1 only with n-hexane, n-heptane or n-octane as alkane substrates (Ehrenreich et al., 2000). Whether or not this pronounced specificity is due to the alkane-activating enzyme or to enzymes for subsequent steps is presently unknown. Denitrifying strain HxN1 converts also butane, pentane and cylcopentane to alkylsuccinates and to fatty acids that are indicative of a further metabolism (Wilkes et al., 2003). The lack of growth on these hydrocarbons suggests that their activation and possibly also their further processing are by-reactions that may occur at rather low rate in comparison to the rates of n-hexane, n-heptane or n-octane utilization.

3.2

Degradation of Alkylsuccinates to Acetyl-CoA

A further degradation of (1-methylalkyl)succinates, upon CoA-ligation of either carboxyl group, cannot proceed through regular b-oxidation and thiolytic cleavage; the formed tertiary carbon (branching sites; > Fig. 3a) exclude oxidation beyond the alcohol level. Different dehydrogenation and hydration reactions may be theoretically formulated to by-pass this problem and allow further degradation via aldol-like cleavage reactions. However, the identification of fatty acids apparently derived from alkanes and isotope labeling studies so far pointed at a carbon skeleton rearrangement (Wilkes et al., 2002). In 4-methyloctanoic acid formed in n-hexane-grown cultures, obviously from the initial product by carboxyl group loss, the single D-atom remaining from added d2-fumarate (see above) in the formed d1-(1-methylpentyl)succinate apparently had been shifted by one position closer to the methyl branch. A plausible explanation was that the D-atom and the carboxyl group (presumably as CoA-thioester) next to the methyl branch exchanged their positions, and that the other carboxyl group was lost by decarboxylation. Such a reaction would be analogous the carbon skeleton rearrangement of succinyl-CoA yielding methylmalonyl-CoA in propionic acid fermentation; methylmalonyl-CoA as a kind of 3-oxoacid, after epimerization of the (R)- to the (S)-form, undergoes decarboxylation (strictly speaking transcarboxylation). Accordingly, the (2-methylhexyl)malonyl-CoA (also termed 2-carboxyl-4-methyloctanoyl-CoA) formed by C-skeleton rearrangement from (1-methylpentyl)succinyl-CoA would decarboxylate (or transcarboxylate) to 4-methyloctanoyl-CoA. The methyl branch in this compound no longer presents an obstacle to b-oxidation. Degradation of hydrocarbon-derived substituted succinates via C-skeleton rearrangement and decarboxylation (or transcarboxylation) was also suggested by studies with other n-alkanes (Davidova et al., 2005; Cravo-Laureau 2005; Callaghan et al., 2006; Kniemeyer et al., 2007) in anaerobes, and even for the degradation of the aromatic hydrocarbon, ethylbenzene, in a sulfate-reducing bacterium (Kniemeyer et al., 2003). However, the suggested pathway has not yet been substantiated by enzymatic measurements, and some details of reactions are still unknown or enigmatic. For instance, metabolite

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

8

analyses not only indicated formation of 4-methyl-2-alkenoic acids from the expected dehydrogenation in b-oxidation, but also formation of 4-methyl-3-alkenoic acids (Wilkes et al., 2002; Cravo-Laureau 2005). The significance of this reaction is unknown.

4

Other Degradation Pathways

The sulfate-reducing bacterium strain Hxd3, tentatively named Desulfococcus (formerly Desulfobacterium) oleovorans, grows on long-chain alkanes such as hexadecane (Aeckersberg et al., 1991). Growth on C-even n-alkanes led to the dominance of C-odd cellular fatty acids, whereas growth on C-odd alkanes resulted mainly in C-even cellular fatty acids (Aeckersberg et al., 1998; So et al., 2003). In contrast, fatty acids remained C-even upon growth with C-even fatty acids (and 1-alkenes) and C-odd upon growth with C-odd fatty acids (and 1-alkenes). Such an alteration of the alkane carbon chain has not been observed with other anaerobic alkane degraders. Initially discussed mechanisms for alkane activation in strain Hxd3 were a terminal (energy-driven) carbonylation of an alkane activated by a transition metal (Aeckersberg et al., 1998) yielding an aldehyde, a reaction known in metal-organic chemistry (Sakakura and Tanaka 1987), and an addition of C-3 (instead of C-2) to fumarate (Rabus et al., 2001), a by-reaction in denitrifying strain HxN1. However, an alkylsuccinate was not detectable in strain Hxd3 (So et al., 2003; Callaghan et al., 2006). Metabolite analyses upon growth with alkanes and 13C-CO2/bicarbonate indicated the presence of fatty acids that were by one C-atom shorter than the parental alkane and carried the 13C-label in the carboxyl group. Such CO2-derived carboxyl label was not incorporated during growth with fatty acids. If a perdeuterated alkane was metabolized, two D-atoms, one at carbon-2 and another at carbon-3, carbon-4 or carbon-5 (position unknown) of the fatty acid were exchanged by H-atoms. It was suggested that the alkane is attacked at C-3 and that a bicarbonate-derived carboxyl group is introduced at this position to yield a 2-ethylcarboxylic acid (> Fig. 5). The D/H-exchange is assumed to occur during activation. The ethyl group of the so far undetected 2-ethylcarboxylic acid was suggested to be removed during further oxidation; this assumption was in accordance with the observation that [1,2–13C2]-hexadecane partly resulted in fatty acids which fully lost the two labels. Anaerobic formation of a 2-ketone as an intermediate from alkanes with subsequent carboxylation at carbon-3 (an otherwise feasible reaction, due to C
H acidity at this position) was excluded. Moreover, the sequenced genome of strain Hxd3 (http://www.jgi. doe.gov/genome-projects/) does not indicate an open reading frame encoding the assumed characteristic glycyl radical protein known from other anaerobic alkane degraders.

5

Terminal Oxidation

Because of their capability for complete oxidation, alkane degraders must further metabolize acetyl-CoA. Only in methanogenic alkane degradation, the alkane-degrading syntrophs (reducing H+-ions to H2 that is consumed by methanogens) may incompletely oxidize their substrate to acetate which is metabolized by methanogens (Zengler et al., 1999) or alternatively by (presumably other) syntrophs (Jones et al., 2008). In denitrifying bacteria, terminal oxidation is generally assumed to proceed via the citric acid cycle, even though this has not been confirmed in alkane-utilizing denitrifiers. A close relative of the alkane degrading strain HxN1, the Betaproteobacterium strain EbN1, degrades alkylbenzenes but not alkanes; the genome of

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Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

. Figure 5 Proposed pathway for the anaerobic degradation of long-chain n-alkanes in the sulfate-reducing bacterium, strain Hxd3. n-Alkanes lead to the dominance of C-odd cellular fatty acids, whereas growth on C-odd alkanes resulted mainly in C-even cellular fatty acids. 13C-labelled bicarbonate (bold) resulted in a labeled carboxyl group of the fatty acid that is by one C-atom shorter than the alkane used for growth. A 13C-label (▲) in C-1 and C-2 of the alkane either disappeared completely, or was conserved in the chain (not shown). These observations lead to the proposed incorporation of a CO2 at C-3, most likely upon a preceding (so far unknown) activation. The proposed branched-chain intermediates have not been detected so far.

strain EbN1 indeed indicated the presence of a citric acid cycle (Rabus et al., 2005), and the same is thus expected for strain HxN1. The most common pathway for acetyl-CoA degradation in sulfate-reducing bacteria is the oxidative Wood-Ljungdahl pathway (C1-pathway), with carbon monoxide dehydrogenase (reverse acetyl-CoA synthetase) as a key enzyme (Schauder et al., 1986). Only Desulfobacter species make use of a modified citric acid cycle (Mo¨ller et al., 1987).

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes

8

It is therefore assumed that also alkane-degrading sulfate-reducing bacteria, which are not closely related to Desulfobacter, make use of the C1-pathway. The key enzyme, carbon monoxide dehydrogenase has indeed been measured in the sulfate-reducing hexadecane degrader, strain Hxd3, while CoA-dependent 2-oxoglutarate dehydrogenase, which is regarded as indicative of a complete citric acid cycle, was absent (Aeckersberg et al., 1991).

6

Regulation

So far investigated, the capacity for the anaerobic degradation of alkanes is regulated, as often in case of enzymes for “fastidious” reactions. A sulfate-reducing strain formed the protein assumed to represent the catalytic subunit of the alkane-activating enzyme upon growth with n-hexadecane, but not with hexadecanoate (Callaghan et al., 2008). Growth of a denitrifying strain with n-hexane started after a lag-phase if the pre-culture was grown with n-hexanoate (Grundmann et al., 2008). Proteins assumed to be part of the alkane-activating enzyme as well as transcripts of the encoding genes were only detectable in n-hexane-grown cells. Hydrocarbon-sensing and gene-regulating systems in anaerobic alkane degraders are still unknown. In the case of denitrifiers that degrade alkylbenzenes, there is evidence for a two-component system for growth with toluene and a one-component system for growth with ethylbenzene (Ku¨hner et al., 2005; and references cited therein).

7

Research Needs

The reactions for alkane activation and consecutive processing that have been formulated on the basis of identified metabolites, isotope labeling studies, proteomic analyses and analogies to the activation of toluene need further corroboration by enzymatic studies. A certain drawback is the relatively elaborate cultivation that is often confronted with cell attachment to floating alkane droplets as well as with slow growth and low cell yields in particular in the case of sulfate-reducing bacteria. Enzymes such as the alkane-activating radical enzyme may not be very stable. The elaboration of modified protocols to improve cultivation as well as stability of the expectedly sensitive enzyme activities (which usually requires ‘‘trial and error’’ approaches) may be desirable. A stable enzymatic activity in vitro would, for instance, allow the study of enzyme specificity by comparison of rates with different alkanes and other hydrocarbons. Can the narrow substrate range of the anaerobic bacterial species with respect to alkanes indeed be explained by a pronounced specificity of their activating enzymes, or are enzymes for subsequent steps also highly specific? A long-term goal, the realization of which is difficult to predict, would be to crystallize and resolve the structure of an alkane-activating enzyme and compare this to the (presently still unknown) structure of a toluene-activating enzyme. The only anaerobic hydrocarbon-activating enzyme with resolved crystal structure is ethylbenzene dehydrogenase from a denitrifying Betaproteobacterium (Kloer et al., 2006), which is not a glycyl radical enzyme (> Chapter 5, Vol. 2, Part 2). Furthermore, alkane activation and further processing also involves a delicate stereochemistry that awaits elucidation (> Fig. 3b). A natural hydrocarbon with essentially unexplored microbiology and biochemistry under anoxic conditions is ethane (C2H6). Ethane possesses only primary carbon atoms.

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Even though an activation of an alkane at a primary carbon atom by a radical-catalyzed addition to fumarate appears in principle possible, as shown by the formation of n-propylsuccinate from propane (Kniemeyer et al., 2007), such a reaction is presumably extremely slow and was supposed to be a side reaction in the anaerobic metabolism of propane. This may explain the extremely slow growth of a sulfate-reducing enrichment culture on ethane (Kniemeyer et al., 2007). The study of the involved microorganisms and metabolites is presumably a tedious and long-term task. Nevertheless, on a global time scale such reactions may be significant, for instance in the alteration of the composition of gas under anoxic conditions in natural reservoirs.

References Aeckersberg F, Bak F, Widdel F (1991) Anaerobic oxidation of saturated hydrocarbons to CO2 by a new type of sulfate-reducing bacterium. Arch Microbiol 156: 5–14. Aeckersberg F, Rainey FA, Widdel F (1998) Growth, natural relationships, cell fatty acids and metabolic adaptation of sulfate-reducing bacteria that utilize long-chain alkanes under anoxic conditions. Arch Microbiol 170: 361–369. Anderson RT, Lovley DR (2000) Hexadecane decay by methanogenesis. Nature 404: 722–723. Becker A, Fritz-Wolf K, Kabsch W, Knappe J, Schultz S, Volker Wagner AF (1999) Structure and mechanism of the glycyl radical enzyme pyruvate formate-lyase. Nat Struct Biol 6: 969–975. Beller HR, Spormann AM (1997) Anaerobic activation of toluene and o-xylene by addition to fumarate in denitrifying strain T. J Bacteriol 179: 670–676. Beller HR, Spormann AM (1998) Analysis of the novel benzylsuccinate synthase reaction for anaerobic toluene activation based on structural studies of the product. J Bacteriol 180: 5454–5457. Biegert T, Fuchs G, Heider J. (1996) Evidence that anaerobic oxidation of toluene in the denitrifying bacterium Thauera aromatica is initiated by formation of benzylsuccinate from toluene and fumarate. Eur J Biochem 238: 661–668. Bregnard TP, Haner A, Hohener P, Zeyer J (1997) Anaerobic degradation of pristane in nitrate-reducing microcosms and enrichment cultures. Appl Environ Microbiol 63: 2077–2081. Buckel W, Golding BT (2006) Radical enzymes in anaerobes. Annu Rev Microbiol 60: 27–49. Callaghan AV, Gieg LM, Kropp KG, Suflita JM, Young LY (2006) Comparison of mechanisms of alkane metabolism under sulfate-reducing conditions among two bacteria isolates and a bacterial consortium. Appl Environ Microbiol 72: 4274–4282. Callaghan AV, Wawrik B, Nı´ Chadhain SM, Young LY, Zylstra GJ (2008) Anaerobic alkane-degrading strain

AK-01 contains two alkylsuccinate synthase genes. Biochem Biophys Res Commun 366: 142–148. Callaghan AV, Tierney M, Phelps CD, Young LY (2009) Anaerobic biodegradation of n-hexadecane by a nitrate-reducing consortium. Appl Environ Microbiol, doi:10.1128/AEM.02491-08. Coschigano PW, Wehrman TS, Young LY (1998) Identification and analysis of genes involved in anaerobic toluene metabolism by strain T1: putative role of a glycine free radical. Appl Environ Microbiol 64: 1650–1656. Cravo-Laureau C, Matheron R, Cayol J-L, Joulian C, Hirschler-Rea A (2004) Desulfatibacillum aliphaticivorans gen. nov., sp. nov., an n-alkane- and n-alkene-degrading, sulphate-reducing bacterium. Int J Syst Evol Microbiol 54: 77–83. Cravo-Laureau C, Grossi V, Raphel D, Matheron R, Hirschler-Rea A (2005) Anaerobic n-alkane metabolism by a sulfate-reducing bacterium, Desulfatibacillum aliphaticivorans strain CV2803T. Appl Environ Microbiol 71: 3458–3467. Davidova IA, Suflita JM (2005) Enrichment and isolation of anaerobic hydrocarbon-degrading bacteria. In: JR (ed.). Leadbetter Methods in enzymology, vol 397. Amsterdam: Elsevier, pp 17–34. Davidova IA, Gieg LM, Nanny M, Kropp KG, Suflita JM (2005) Stable isotope studies of n-alkane metabolism by a sulfate-reducing bacterial enrichment culture. Appl Environ Microbiol 71: 8174–8182. Eastcott L, Shiu WY, Mackay D (1988) Environmentally relevant physical-chemical properties of hydrocarbons: a review of data and developments of simple correlations. Oil Chem Pollut 4: 191–216. Ehrenreich P, Behrends A, Harder J, Widdel F (2000) Anaerobic oxidation of alkanes by newly isolated denitrifying bacteria. Arch Microbiol 173: 58–64. Frey PA, Hegeman AD, Ruzicka FJ (2008) The radical SAM superfamily. Crit Rev Biochem Mol Biol 43, 63–88. Gieg LM, Suflita JM (2002) Detection of anaerobic metabolites of saturated and aromatic hydrocarbons in

Biochemistry of the Anaerobic Degradation of Non-Methane Alkanes petroleum-contaminated aquifers. Environ Sci Technol 36: 3755–3762. Grundmann O, Behrends A, Rabus R, Amann J, Halder T, Heider J, Widdel F (2008) Genes encoding the candidate enzyme for anaerobic activation of n-alkanes in the denitrifying bacterium, strain HxN1. Environ Microbiol 10: 376–385. Jones DM, Head IM, Gray ND, Adams JJ, Rowan AK, Aitken CM, Bennett B, Huang H, Brown A, Bowler BF, Oldenburg T, Erdmann M, Larter SR (2008) Crude-oil biodegradation via methanogenesis in subsurface petroleum reservoirs. Nature 451: 176–180. Jordan A, Reichard P (1998) Ribonucleotide reductases. Annu Rev Biochem 67: 71–98. Kloer DP, Hagel C, Heider J, Schulz GE (2006) Crystal structure of ethylbenzene dehydrogenase from Aromatoleum aromaticum. Structure 14: 1377–1388. Knappe J, Neugebauer FA, Blaschkowski HP, Ga¨nzler M (1984) Post-translational activation introduces a free radical into pyruvate formate-lyase. Proc Natl Acad Sci USA 81: 1332–1335. Kniemeyer O, Fischer T, Wilkes H, Glo¨ckner FO, Widdel F (2003) Anaerobic degradation of ethylbenzene by a new type of marine sulfate-reducing bacterium. Appl Environ Microbiol 69: 760–768. Kniemeyer O, Musat F, Sievert S, Knittel K, Wilkes H, Blumenberg M, Michaelis W, Classen A, Bolm C, Joye S, Widdel F (2007) Anaerobic oxidation of short-chain hydrocarbons by marine sulphatereducing bacteria. Nature 449: 898–901. Krieger CJ, Roseboom W, Albracht SP, Spormann AM (2001) A stable organic free radical in anaerobic benzylsuccinate synthase of Azoarcus sp. strain T. J Biol Chem 276: 12924–12927. Kropp KG, Davidova IA, Suflita JM (2000) Anaerobic oxidation of n-dodecane by an addition reaction in a sulfate-reducing bacterial enrichment culture. Appl Environ Microbiol 66: 5393–5398. Ku¨hner S, Wo¨hlbrand L, Fritz I, Wruck W, Hultschig C, Hufnagel P, Kube M, Reinhardt R, Rabus R (2005) Substrate-dependent regulation of anaerobic degradation pathways for toluene and ethylbenzene in a denitrifying bacterium, strain EbN1. J Bacteriol 187: 1493–1503. Leuthner B, Leutwein C, Schulz H, Ho¨rth P, Haehnel W, Schiltz E, Scha¨gger H, Heider J (1998) Biochemical and genetic characterization of benzylsuccinate synthase from Thauera aromatica: a new glycyl radical enzyme catalysing the first step in anaerobic toluene metabolism. Mol Microbiol 28: 615–628. Leutwein C, Heider J (1999) Anaerobic toluene-catabolic pathway in denitrifying Thauera aromatica: activation and beta-oxidation of the first intermediate, (R)-(+)-benzylsuccinate. Microbiology 145: 3265–3271.

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Mo¨ller D, Schauder R, Fuchs G, Thauer RK (1987) Acetate oxidation to CO2 via a citric acid cycle involving an ATP-citrate lyase: a mechanism for the synthesis of ATP via substrate level phosphorylation in Desulfobacter postgatei growing on acetate and sulfate. Arch Microbiol 148: 202–207. Musat F, Galushko A, Jacob J, Widdel F, Kube M, Reinhardt R, Wilkes H, Schink B, Rabus R (2009) Anaerobic degradation of naphthalene and 2-methylnaphthalene by strains of marine sulfatereducing bacteria. Environ Microbiol 11: 209–219. O’Brien JR, Raynaud C, Croux C, Girbal L, Soucaille P, Lanzilotta WN (2004) Insight into the mechanism of the B12-independent glycerol dehydratase from Clostridium butyricum: preliminary biochemical and structural characterization. Biochemistry 43: 4635–4645. Qiao C, Marsh ENG (2005) Mechanism of benzylsuccinate synthase: stereochemistry of toluene addition to fumarate and maleate. J Am Chem Soc 127: 8608–8609. Rabus R, Wilkes H, Behrends A, Armstroff A, Fischer T, Pierik AJ, Widdel F (2001) Anaerobic initial reaction of n-alkanes: evidence for (1-methylpentyl)succinate as initial product and for involvement of an organic radical in the metabolism of n-hexane in a denitrifying bacterium. J Bacteriol 183: 1707–1715. Rabus R, Kube M, Heider J, Beck A, Heitmann K, Widdel F, Reinhardt R (2005) The genome sequence of an anaerobic aromatic-degrading denitrifying bacterium, strain EbN1. Arch Microbiol 183: 27–36. Rueter P, Rabus R, Wilkes H, Aeckersberg F, Rainey FA, Jannasch HW, Widdel F (1994) Anaerobic oxidation of hydrocarbons in crude oil by new types of sulphate-reducing bacteria. Nature 372: 455–458. Sakakura T, Tanaka M (1987) Efficient catalytic C
H activation of alkanes: regioselective carbonylation of the terminal methyl group of pentane by RhCl(CO)(PMe3)2. J Chem Soc Chem Commun (no volume number): 758-759. Sawers G, Hesslinger C, Muller N, Kaiser M (1998) The glycyl radical enzyme TdcE can replace pyruvate formate-lyase in glucose fermentation. J Bacteriol 180: 3509–3516. Schauder R, Eikmanns B, Thauer RK, Widdel F, Fuchs G (1986) Acetate oxidation to CO2 in anaerobic bacteria via a novel pathway not involving reactions of the citric acid cycle. Arch Microbiol 145: 162–172. Selmer T, Andrei PI (2001) p-Hydroxyphenylacetate decarboxylase from Clostridium difficile. A novel glycyl radical enzyme catalysing the formation of p-cresol. Eur J Biochem 268: 1363–1372. Shinoda Y, Akagi J, Uchihashi Y, Hiraishi A, Yukawa H, Yurimoto H, Sakai Y, Kato N (2005) Anaerobic degradation of aromatic compounds by Magnetospirillum

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strains: isolation and degradation genes. Biosci Biotechnol Biochem 69: 1483–1491. So CM, Young LY (1999) Isolation and characterization of a sulfate-reducing bacterium that anaerobically degrades alkanes. Appl Environ Microbiol 65: 2969–2976. So CM, Phelps CD, Young LY (2003) Anaerobic transformation of alkanes to fatty acids by a sulfate-reducing bacterium, strain Hxd3. Appl Environ Microbiol 69: 3892–3900. Townsend GT, Prince RC, Suflita JM (2004) Anaerobic biodegradation of alicyclic constituents of gasoline and natural gas condensate by bacteria from an anoxic aquifer. FEMS Microbiol Ecol 49: 129–135. Verfu¨rth K, Pierik AJ, Leutwein C, Zorn S, Heider J (2004) Substrate specificities and electron paramagnetic resonance properties of benzylsuccinate synthases in anaerobic toluene and m-xylene metabolism. Arch Microbiol 181: 155–162. Wagner AF, Frey M, Neugebauer FA, Scha¨fer W, Knappe J (1992) The free radical in pyruvate formate-lyase is

located on glycine-734. Proc Natl Acad Sci USA 89: 996–1000. Wilhelm E, Battino R, Wilcock RJ (1977) Low-pressure solubility of gases in liquid water. Chem Rev 77: 219–262. Wilkes H, Ku¨hner S, Bolm C, Fischer T, Classen A, Widdel F, Rabus R (2003) Formation of n-alkaneand cycloalkane-derived organic acids during anaerobic growth of a denitrifying bacterium with crude oil. Organ Geochem 34: 1313–1323. 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. Zengler K, Richnow HH, Rosello´-Mora R, Michaelis W, Widdel F (1999) Methane formation from longchain alkanes by anaerobic microorganisms. Nature 401: 266–269.

9 Anaerobic Degradation of Aromatic Hydrocarbons M. Tierney . L. Y. Young Biotechnology Center for Agriculture and the Environment, Rutgers University, New Brunswick, NJ, USA [email protected] 1 1.1 1.2

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 926 Anaerobic Mechanisms: Fumarate Addition Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . 926 Anaerobic Mechanisms: Carboxylation Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 926

2 2.1 2.2 2.3 2.4 2.4.1 2.4.2 2.4.3

Polycylic Aromatic Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 927 General Description . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 927 Anaerobic PAH Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 927 Pure Cultures of Anaerobic PAH Degraders . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 928 Anaerobic PAH Degradation Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 928 Naphthalene Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 928 2-Methylnaphthalene Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 929 Phenanthrene Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 929

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 930

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Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 931

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_65, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Many anaerobic prokaryotes are capable of mineralizing petroleum-derived hydrocarbons, including alkanes and aromatic compounds. The rate of microbial growth, cell yield, and the amount of energy released as a result of metabolism depends on the catabolic pathway and terminal electron acceptor utilized. Nitrate-reducing, denitrifying, iron-reducing, sulfatereducing, methanogenic, and anoxygenic phototrophic organisms have been linked to aromatic degradation. Of these potential electron acceptors, NO3 is the most energetic for microbes, followed by Fe3+, SO4 , and CO2.

1

Introduction

1.1

Anaerobic Mechanisms: Fumarate Addition Reactions

Radical enzymes play an important role in the degradation of aromatic and alkyl hydrocarbons; they aid in reducing the stability of the aromatic nucleus (Gibson and Harwood, 2002). Initially, insights into anaerobic degradation of toluene were reported in Evans et al. (1992). Evans isolated an anaerobic, denitrifying bacterium, designated T1, and found evidence of benzylsuccinate production in toluene grown cultures (Evans, 1991, 1992). Early studies with T1 were instrumental in the genetic characterization of anaerobic toluene degradation (Coschigano, 2000; Coschigano et al.,1998; Coschigano and Young, 1997). In anaerobic toluene oxidation, benzylsuccinate synthase (bss) facilitates the addition of fumarate to the methyl group of toluene (Leuthner and Heider, 2000). The enzyme abstracts a hydrogen atom from the methyl group of toluene and the methyl group is then added to the double bond of fumarate. The resulting intermediate, benzylsuccinate, retains the H in succinyl moiety. Benzoyl-CoA, a common monoaromatic breakdown product, is generated after benzylsuccinate undergoes ß-oxidation (Beller and Spormann, 1998; Gibson and Harwood, 2002; Leuthner and Heider, 2000). This degradation mechanism is conserved among various types of anaerobic microorganisms utilizing toluene and other substituted monaromatic contaminants, including denitrifiers, sulfate-reducers, methanogens, and anoxygenic phototrophs (Beller and Edwards, 2000; Kniemeyer et al., 2003; Krieger et al., 1999; Zengler et al., 1999). Methylated compounds including m and p-cresol, 2-methylnaphthalene, and alkanes are also reportedly degraded using this catabolic strategy (Annweiler et al., 2000; ASM News, 1999; Callaghan et al. 2006; 2008; Gibson and Harwood, 2002; Meckenstock et al., 2004). Recently, Safinowski and Meckenstock (2006) reported finding a fumarate addition intermediate in sulfidogenic, naphthalene-fed enrichment cultures (Safinowski and Meckenstock, 2006).

1.2

Anaerobic Mechanisms: Carboxylation Reactions

While many bacteria utilize a radical enzyme facilitated fumarate addition strategy to anaerobically degrade a wide range of compounds, carboxylation is also a catabolic mechanism shared among anaerobes. The addition of CO2 or bicarbonate to an organic acceptor is a very relevant biological reaction necessary for a range of cellular functions. Many compounds, including alkanes, can be microbially activated through carboxylation reactions for downstream catabolic reactions (Callaghan et al., 2006; Rabus et al.,2001; So et al., 2003).

Anaerobic Degradation of Aromatic Hydrocarbons

9

Monoaromatic compounds can also be activated by carboxylation. For example, phenol, a hydroxyl substituted benzene ring, is converted to phenylphosphate by phenylphosphate synthase. Phenylphosphate carboxylase, an oxygen-sensitive enzyme that uses CO2 as a substrate, converts phenylphosphate to 4-hydroxy-benzoate. 4-hydroxy-benzoate is then transformed to 4-hydroxy-benzoyl CoA, which is further metabolized to benzoyl-CoA (Breinig et al., 2000; Lack et al., 1991; Schuhle and Fuchs, 2004). Polycyclic aromatic hydrocarbons (PAHs) can also be activated by carboxylation (Zhang et al., 2000; Zhang and Young, 1997). Studies by Zhang et al. (1997) and Davidova et al. (2007) provide evidence that two and three-ringed PAHs are carboxylated as an initial activation mechanism (Davidova et al.,2007; Zhang and Young, 1997).

2

Polycylic Aromatic Hydrocarbons

2.1

General Description

Aromatic hydrocarbons are found in gasoline and are manufactured as industrial solvents. Polycyclic aromatic hydrocarbons (PAHs) are formed during incomplete combustion of organic compounds like those found in coal, oil, and gas. These compounds are comprised of fused aromatic rings, which make them very stable due to the resonance energy of the ring structure. PAHs are EPA priority pollutants and are regulated as toxic waste under Resource Conservation and Recovery Act (RCRA). PAH contamination is widespread in the environment due to pyrolytic and industrial activity (Bamforth and Singleton, 2005; Samanta and Jain, 2000). PAHs have carcinogenic, mutagenic, and toxic properties (Huntley et al., 1993; Weissenfels et al., 1992). PAHs are only carcinogenic in mammals and higher animals once they enter and have been activated by the cell. Metabolic activation by cytochrome p450 in mammalian cells produces intermediates more harmful than the parent compound, presenting toxic and mutagenic risks to the organism (Bamforth and Singleton, 2005; Zylstra et al.1997). The properties of PAHs make them amenable to biological and physical transformations. PAHs have low aqueous solubilities, high octanol-water partition coefficients, and vapor pressures (Huntley et al., 1993). As a result of these characteristics, PAHs have a high affinity for particulates. PAHs are transported to sediments through atmospheric deposition, industrial effluents, and wastewater treatment plant discharges (Bamforth and Singleton, 2005). PAH removal in sediments is mainly by microorganisms, but PAHs may also undergo physical processes including volatilization, photolysis, and chemical oxidation (Weissenfels et al., 1992).

2.2

Anaerobic PAH Degradation

Since PAHs are chemically stable in the environment due to the resonance energy of the aromatic ring system, microbial activation and metabolism is an essential component of PAH reduction in sediments and soils. Biodegradation of PAHs has been shown to occur under various redox conditions, indicating that a range of microorganisms have the metabolic capability to degrade these pollutants (Bedessem et al., 1997; Chang et al., 2002; Coates et al., 1996; 1997; Eriksson et al., 2003; McNally et al., 1999; Meckenstock et al., 2004;

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Ramsay et al., 2003; Rockne et al., 2000; Rockne and Strand, 2000; Sharak Genther et al.,1997; Zhang and Young, 1997). A general trend has been observed under anaerobic conditions that lower molecular weight PAHs are less recalcitrant than higher molecular weight PAHs (Johnson and Ghosh, 1998). PAHs are readily degraded under anaerobic conditions. It has been reported that anaerobic PAH degradation rates are fastest under sulfidogenic conditions, followed by methanogenic conditions, and finally nitrate-reducing conditions (Chang et al., 2002). Naphthalene and anthracene are degraded to some extent under iron-reducing conditions (Ramsay et al., 2003). Naphthalene, methylnaphthalene, phenanthrene, and fluorene were reportedly oxidized in harbor sediments under sulfate-reducing conditions (Coates et al., 1996, 1997). Naphthalene has also been shown to be degraded in enrichment culture under sulfidogenic, methanogenic, and nitrate-reducing conditions (Bedessem et al., 1997; Meckenstock et al., 2000; Rockne and Strand, 2000; Sharak Genther et al., 1997). 2-methylnaphthalene and the tricyclic PAH anthraquinone are biodegraded under methanogenic conditions (Sharak Genther et al., 1997). Phenanthrene and pyrene have been shown to be transformed under denitrifying conditions (McNally et al., 1999; Rockne and Strand, 2000). Acenaphthene, 3-methylnaphthalene, fluorene, and phenanthrene are reportedly degraded under nitrate-reducing conditions (Eriksson et al., 2003; Mihelcic and Luthy, 1988).

2.3

Pure Cultures of Anaerobic PAH Degraders

Naphthalene is the only PAH shown to be anaerobically degraded by pure-cultures. Degradation has been established under nitrate-reducing and sulfidogenic conditions. Three anaerobic isolates reportedly utilize PAHs: a sulfate-reducer, designated NaphS2, a nitrate-reducer, and a denitrifier have been shown to degrade naphthalene anaerobically (Galushko et al., 1999; Rockne et al., 2000). The pure culture NaphS2 has been shown to use 2-naphthoic acid, an intermediate of naphthalene metabolism, as a growth substrate (Galushko et al., 1999). Rockne et al. (2000) isolated two nitrate-dependent naphthalene degraders. Phylogenetic analyses reveal that these isolates are most closely related to Pseudomonas stutzeri and Vibrio pelagius (Rockne et al., 2000). Although these cultures are available as pure strains, no pathway or genetic analyses have been reported.

2.4

Anaerobic PAH Degradation Pathways

2.4.1

Naphthalene Degradation

Three mechanisms have been proposed for naphthalene activation by sulfidogenic bacteria: hydroxylation, carboxylation and methylation (Bedessem et al.1997; Safinowski and Meckenstock, 2006; Zhang and Young, 1997). In 1997, Bedessem et al. proposed that hydroxylation is the first step in anaerobic naphthalene degradation (Bedessem et al., 1997). Also in 1997, Zhang and Young proposed carboxylation as the initial reaction in naphthalene catabolism (Zhang and Young, 1997). Isotope labeling experiments show that carbon derived from 13 C-labeled bicarbonate is directly incorporated onto the aromatic nucleus. They then proposed carboxylation as a mechanism to activate naphthalene and form 2-naphthoic acid (Zhang and Young, 1997).

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Safinowski and Meckenstock (2006) found evidence of a degradation strategy analogous to anaerobic monoaromatic hydrocarbon metabolism (Safinowski and Meckenstock, 2006). They propose that naphthalene is first methylated to facilitate a fumarate addition reaction. In accordance with data published by Zhang and Young (1997), methylation may be facilitated by CO-dehydrogenase, an enzyme associated with acetate oxidation, which can theoretically generate a methyl group from bicarbonate. After methylation, a fumarate moiety is added at the methyl group, generating the intermediate naphthyl-2-methyl-succinic acid. Naphthyl-2-methyl-succinic acid is then oxidized to naphthyl-2-methylene-succinic acid (Safinowski and Meckenstock, 2006). A series of proposed oxidation reactions ultimately results in the formation of 2-naphthoic acid, a central metabolite of anaerobic PAH degradation (McNally et al., 1999; Safinowski and Meckenstock, 2006). Zhang and Young (1997) demonstrated that naphthalene transformed to 2-naphthoic acid. In a later report, Zhang et al. (2000) identified a lower pathway of naphthalene degradation (Zhang et al.2000). The ring system is sequentially reduced, generating the experimentally resolved metabolites tetrahydro-2-naphthoic acid, hexahydro-2-naphthoic acid, and deca-hydro-2-naphthoic acid (Aitken et al., 2004; Annweiler et al., 2002; Zhang et al., 2000). Refer to > Fig. 1.

2.4.2

2-Methylnaphthalene Degradation

In 2-methylnaphthalene metabolism, a mechanism similar to that seen in anaerobic toluene degradation appears to be utilized (Annweiler et al., 2000; Meckenstock et al., 2004). A fumarate moiety is added at the methyl group, resulting in the intermediate naphthyl2-methyl-succinic acid. Naphthyl-2-methyl-succinic acid is then oxidized to naphthyl-2methylene-succinic acid (Annweiler et al., 2000). A series of reactions ultimately results in the formation of 2-naphthoic acid, a proposed central metabolite in PAH degradation (Meckenstock et al., 2004). Further metabolism of 2-naphthoic acid proceeds in the same manner as described for naphthalene (Zhang et al., 2000).

2.4.3

Phenanthrene Degradation

Studies by Zhang and Young (1997) of a sulfate-reducing enrichment culture grown on phenanthrene supplemented with 13C-labeled bicarbonate generated stable isotope labeled phenanthroic acid. This demonstrated that the carbon of carboxyl group originated from 13 C-labeled bicarbonate, suggesting that carboxylation was the activation mechanism (Zhang and Young, 1997). Data obtained by Davidova et al. (2007) derived from a sulfidogenic enrichment culture also indicates uptake of stable isotope from 13C-labeled bicarbonate, further supporting the observations of Zhang and Young (Davidova et al., 2007). Phenanthrene-carboxaldehyde has also been found in nitrate-reducing phenanthrenefed cultures. This suggests that similar mechanisms to activate phenanthrene may be utilized by organisms that thrive under different redox conditions (Eriksson et al., 2003; Zhang and Young, 1997).

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. Figure 1 Anaerobic naphthalene degradation pathway. Proposed anaerobic naphthalene degradation pathways by sulfate-reducing bacteria. Structures represent experimentally resolved metabolites. This figure is adapted from Zhang and Young (1997), Zhang et al. (2000), Bedessem et al. (1997), and Safinowski and Meckenstock (2006).

3

Research Needs

The knowledge that biodegradation of monocyclic, polycylic and alkane hydrocarbons takes place in the absence of oxygen opens up a large number of questions still to be answered. Currently work on the functional genes has been limited to the bss genes and their analogues operating in the fumarate addition mechanism. Evidence suggests, however, that other

Anaerobic Degradation of Aromatic Hydrocarbons

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mechanisms have evolved to allow microorganisms to access these rich carbon sources. For example, although stable isotope data support direct carboxylation of naphthalene, description of the chemical mechanism and the genes involved still remain to be described. Detailed mechanistic and genetic work could lead to new biosynthetic pathways for chemicals and well as improving bioremediation of contaminated environments. Additional research is also needed to understand first, how widespread these mechanisms are distributed in the anaerobic community, and second, the impact of these microbes in the environment on the rate and extent of hydrocarbon degradation in soils, sediments and other anoxic habitats.

4

Conclusions

Increasing evidence indicates that bacteria that thrive in the absence of oxygen use similar catabolic strategies to biodegrade a wide variety of environmental contaminants. Fumarate addition and carboxylation reactions are instrumental in this degradation; these reactions facilitate the activation and subsequent mineralization of alkanes, as well as monocyclic and polycyclic aromatic compounds, by a growing number of anaerobic microbes.

Acknowledgments The authors thankfully acknowledge the research contributions of Kelly Hess and the insight of members of the Young laboratory. We are especially grateful for the guidance of Lora McGuinness, Dr. Lee Kerkhof, Dr. David Scala, and Erin Gallagher. This work was supported by NSF CHE 0221978, the Cook College General Honors program, the Biotechnology Summer Internship program, and the Aresty Research Center for Undergraduates.

References Alexander M (1995) How toxic are chemicals in soil? Environ Sci and Technol 29: 2713–2717. Aitken CM, Jones DM, Larter SR (2004) Anaerobic hydrocarbon biodegradation in deep subsurface oil resevoirs. Nature 431: 291–294. Ambrosoli R, et al. (2005) Anaerobic PAH degradation in soil by a mixed bacterial consortium under denitrifying conditions. Chemosphere 60: 1231–1236. Annweiler E, et al. (2000) Anaerobic degradation of 2-methylnaphthalene by a sulfate-reducing enrichment culture. Appl Environ Microbiol 66: 5329–5333. Annweiler E, Mrichaelis W, Meckenstock RU (2002) Identical ring cleavage products during anaerobic degradation of naphthalene, 2-methylnaphthalene, and tetralin indicate a new metabolic pathway. Appl Environ Microbiol 68: 852–858. ASM News (1999) Microbiology’s fifty most significant events during the past 125 years. Poster Supplement 65 65(3).

Atlas RM (1997) Principles of Microbiology. Boston: WCB Publishers. Baker GC, Smith JJ, Cowan DA (2003) Review and reanalysis of domain-specific 16S primers. J Microbiol Meth 55: 541–555. Bamforth SM, Singleton (2005) Bioremediation of polycyclic aromatic hydrocarbons: current knowledge and future directions. J Chem Technol Biotechnol 80: 723–736. Bedessem ME, Swoboda-Colberg NG, Colberg PJS (1997) Naphthalene mineralization coupled to sulfate reduction in aquifer-derived enrichments. FEMS Microbiol Lett 152: 213–218. Beller HR, Edwards ER (2000) Anaerobic toluene activation by benzylsuccinate synthase in a highly enriched methanogenic culture. Appl Environ Microbiol 66: 5503–5505. Beller HR, Spormann AM (1998) Analysis of the novel benzylsuccinate synthase reaction for anaerobic

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toluene activation based on structural studies of the product. J Bacteriol 180: 5454–5457. Bossert ID, Rivera MD, Young LY (1986) p-Cresol biodegradation under denitrifying conditions: Isolation of a bacterial culture. FEMS Microbiol Ecol 385: 313–319. Breinig S, Schiltz E, Fuchs G (2000) Genes involved in anaerobic metabolism of phenol in the bacterium Thauera aromatica. J Bacteriol 182: 5849–5863. Callaghan AV, et al. (2006) Comparison of mechanisms of alkane metabolism under sulfate-reducing conditions among two bacterial isolates and a bacterial consortium. Appl Environ Microbiol 72: 4274–4282. Callaghan AV, et al. (2008) Anaerobic alkane-degrading strain AK-01 contains two alkylsuccinate synthase genes. Biochem Biophys Res Commun 366: 142–148. Chakraborty R, Coates JD (2004) Anaerobic degradation of monoaromatic hydrocarbons. Appl Microbiol Biotechnol 64: 437–446. Chang BV, Shiung LC, Yuan SY (2002) Anaerobic biodegradation of polycyclic aromatic hydrocarbon in soil. Chemosphere 48: 717–724. Chang W, Um Y, Pulliam Holoman TR (2006) Polycyclic aromatic hydrocarbon (PAH) degradation coupled to methanogenesis. Biotechol Letters 28: 425–430. Coates JD, Anderson RT, Lovely DR (1996) Oxidation of polycyclic aromatic hydrocarbons under sulfatereducing conditions. Appl Environ Microbiol 62: 1099–1101. Coates JD, et al. (1997) Anaerobic degradation of polycyclic aromatic hydrocarbons and alkanes in petroleum-contaminated marine harbor sediments. Appl Environ Microbiol 63: 3589–3593. Coschigano PW (2000) Transcriptional analysis of the tutE tutFDGH gene cluster from Thauera aromatica strain T1. Appl Environ Microbiol 66: 1147–1151. Coschigano PW, Wehrman TS, Young LY (1998) Identification and analysis of genes involved in anaerobic toluene metabolism by strain T1: putative role of a glycine free radical. Appl Environ Microbiol 64: 1650–1656. Coschigano PW, Young LY (1997) Identification and sequence analysis of two regulatory genes involved in anaerobic toluene metabolism by strain T1. Appl Environ Microbiol 63: 652–660. Davidova IA, et al. (2007) Anaerobic phenanthrene mineralization by a carboxylating sulfate-reducing bacterial enrichment. ISME J 1: 436–442. Eriksson M, et al. (2003) Degradation of polycyclic aromatic hydrocarbons at low temperature under aerobic and nitrate-reducing conditions in enrichment cultures from northern soils. Appl Environ Microbiol 69: 275–284. Evans PJ, et al. (1991) Anaerobic degradation of toluene by a denitrifying bacterium. Appl Environ Microbiol 57: 1139–1145.

Evans PJ, et al. (1992) Metabolites formed during anaerobic transformation of toluene and o-xylene and their proposed relationship to the initial steps of toluene mineralization. Appl Environ Microbiol 58: 496–501. Forney LJ, Zhou X, Brown CJ (2004) Molecular microbial ecology: land of the one-eyed king. Curr Opin Microbiol 7: 210–220. Gallagher E, et al. (2005) 13C-Carrier DNA shortens the incubation time needed to detect benzoate-utilizing denitrifying bacteria by stable-isotope probing. Appl Environ Microbiol 71: 5192–5196. Galushko A, et al. (1999) Anaerobic degradation of naphthalene by a pure culture of a novel type of marine sulphate-reducing bacterium. Environ Microbiol 1: 415–520. Gibson J, Harwood CS (2002) Metabolic diversity in aromatic compound utilization by anaerobic microbes. Annual Rev Microbiol 56: 345–369. Ginige MP, et al. (2004) Use of stable-isotope probing, full-cycle rRNA analysis, and fluorescence in situ hybridization-microautoradiography to study a methanol-fed denitrifying microbial community. Appl Environ Microbiol 70: 588–596. Glazer AN, Nikaido H (1995) Microbial Biotechnology: Fundamentals of Applied Microbiology. New York: W. H. Freeman and Company. Grosser RJ, et al. (2000) Effect of model sorptive phases on phenanthrene biodegradation: different enrichment conditions influence bioavailability and selection of phenanthrene-degrading isolates. Appl Environ Microbiol 66: 2695–2702. Huntley SL, et al. (1993) Distribution of polycyclic aromatic hydrocarbons (PAHs) in three northern New Jersey waterways. Bull Environ Contam Toxicol 51: 865–872. Jeon CO, et al. (2003) Discovery of a bacterium, with distinctive dioxygenase, that is responsible for in situ biodegradation in contaminated sediment. PNAS 100: 13591–13596. Johnson K, Ghosh S (1998) Feasibility of anaerobic biodegradation of PAHs in dredged river sediments. Wat Sci Tech 38: 41–48. Kanagawa T (2003) Bias and artifacts in multitemplate polymerase chain reactions (PCR). J Biosci Bioeng 96: 317–323. Kasai Y, et al. (2006) RNA-based stable isotope probing and isolation of anaerobic benzene-degrading bacteria from gasoline-contaminated groundwater. Appl Environ Microbiol 72: 3586–3592. Kelsey JW, Kottler BD, Alexander M (1997) Selective chemical extractants to predict bioavailability of soil-aged organic chemicals. Environ Sci and Technol 31: 214–217. Kniemeyer O, et al. (2003) Anaerobic degradation of ethylbenzene by a new type of marine sulfatereducing bacterium strain EBS7. Appl Environ Microbiol 69: 760–768.

Anaerobic Degradation of Aromatic Hydrocarbons Krieger CJ, et al. (1999) Initial reactions in anaerobic oxidation of m-xylene by the denitrifying bacterium Azoarcus sp. strain T. J Bacteriol 181: 6403–6410. Lack A, et al. (1991) Catalytic properties of phenol carboxylase. In vitro study of CO2: 4-hydroxybenzoate isotope exchange reaction. Eur J Biochem 197: 473–479. Leuthner B, Heider J (2000) Anaerobic toluene catabolism of Thauera aromatica: the bbs operon codes for enzymes of beta oxidation of the intermediate benzylsuccinate. J Bacteriol 182: 272–277. Liu W, et al. (1997) Characterization of microbial diversity by determining terminal restriction fragment length polymorphisms of genes encoding 16S rRNA. Appl Environ Microbiol 63: 4516–4522. Madsen EL (2006) The use of stable-isotope probing techniques in bioreactor and field studies on bioremediation. Curr Opin Biotech 17: 92–97. Manefield M, et al. (2002) RNA stable isotope probing, a novel means of linking microbial community function to phylogeny. Appl Environ Microbiol 68: 5367–5373. Marsh TL, et al. (2000) Terminal restriction fragment polymorphism analysis program, a web-based research tool for microbial community analysis. Appl Environ Microbiol 66: 3616–3620. McNally DL, Mihelcic JR, Leuking DR (1999) Biodegradation of mixtures of polycyclic aromatic hydrocarbons under aerobic and nitrate-reducing conditions. Chemosphere 38: 1313–1321. Meckenstock RU, et al. (2000) Anaerobic naphthalene degradation by a sulfate-reducing enrichment culture. Appl Environ Microbiol 66: 2743–2747. Meckenstock RU, Safinowski M, Griebler C (2004) Anaerobic degradation of polycyclic aromatic hydrocarbons. FEMS Microbiol Ecol 49: 27–36. Mihelcic JR, Luthy RG (1988) Microbial degradation of acenaphthene and naphthalene under denitrification conditions in soil-water systems. Appl Environ Microbiol 54: 1188–1198. Morris SA, Radajewski S, Willison TW, Murrell JC (2002) Identification of the functionally active methanotroph population in a peat soil microcosm by stable-isotope probing. Appl Environ Microbiol 68: 1446–1453. Nam K, Alexander M (1998) Role of nanoporosity and hydrophobicity in sequestration and bioavailability: tests of model solids. Environ Sci Technol 32: 71–74. Padmanabhan P, et al. (2003) Respiration of 13C-labeled substrates added to soil in the field and subsequent 16S rRNA gene analysis of 13C-labeled soil DNA. Appl Environ Microbiol 69: 1614–1622. Perez-Jimenez J, Zanaroli JG, Fava F, Young LY (2004) Molecular characterization of indigenous microbiota in PCB-containing microcosms and sediments

9

from the Venice Lagoon (Italy). New Orleans, LA: ASM General Meeting. Phelps CD, Young LY (2001) Biodegradation of BTEX under anaerobic conditions: A Review. Adv Agronom 70: 329–357. Rabus R, et al. (2001) Anaerobic initial reaction of n-alkanes in a denitrifying bacterium: evidence for (1-methylpentyl) succinate as initial product for involvement of an organic radical in n-hexane metabolism. J Bacteriol 183: 1707–1715. Radajewski S, et al. (2000) Stable-isotope probing as a tool in microbial ecology. Nature 403: 646–649. Ramsay JA, et al. (2003) Naphthalene and anthracene mineralization linked to oxygen, nitrate, Fe(III) and sulphate reduction in a mixed microbial population. Biodegradation 14: 321–329. Rockne KJ, et al. (2000) Anaerobic naphthalene degradation by microbial pure cultures under nitratereducing conditions. Appl Environ Microbiol 66: 1595–1601. Rockne KJ, Strand SE (2000) Anaerobic biodegradation of naphthalene, phenanthrene, and biphenyl by a denitrifying enrichment culture. Wat Res 35: 291–299. Safinowski M, Meckenstock RU (2006) Methylation is the initial reaction in anaerobic naphthalene degradation by a sulfate-reducing enrichment culture. Environ Microbiol 8: 347–352. Samanta SK, Jain RK (2000) Evidence for plasmidmediated chemotaxis of Pseudomonas putida towards naphthalene and salicylate. Can J Microbiol 46: 1–6. Scala DJ, Kerkhof LJ (2000) Horizontal heterogeneity of denitrifying bacterial communities in marine sediments by terminal restriction fragment length polymorphism analysis. Appl Environ Microbiol 66: 1980–1986. Schuhle K, Fuchs G (2004) Phenylphosphate carboxylase: a new C-C lyase involved in anaerobic phenol metabolism in Thauera aromatica. J Bacteriol 186: 4556–4567. Sharak Genther BR, et al. (1997) Persistence of polycyclic aromatic hydrocarbon components of creosote under anaerobic enrichment conditions. Arch Environ Contam Toxicol 32: 99–105. So CM, Phelps CD, Young LY (2003) Anaerobic transformation of alkanes to fatty acids by a sulfate-reducing bacterium, strain HXD3. Appl Environ Microbiol 69: 3892–3900. Tang J, et al. (1998) Combined effects of sequestration and bioremediation in reducing the availability of polycyclic aromatic hydrocarbons in soil. Environ Sci Technol 32: 3586–3590. Wackett LP (2004) Stable isotope probing in biodegradation research. Trends Biotech 22: 153–154.

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Weissenfels WD, Klewer HJ, Langhoff S (1992) Adsorption of polycyclic aromatic hydrocarbons (PAHs) by soil particles: influence on biodegradability and biotoxicity. Appl Microbiol Biotech 36: 689–686. Woese CR (1987) Bacterial evolution. Microbiol Rev 51: 221–271. Zengler K, et al. (1999) Phototrophic utilization of toluene under anoxic conditions by a new strain of Blastochloris sulfoviridus. Arch Microbiol 172: 204–212. Zhang X, Sullivan ER, Young LY (2000) Evidence for aromatic ring reduction in the biodegradation

pathway of carboxylated naphthalene by a sulfatereducing consortium. Biodegradation 11: 117–124. Zhang X, 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. Zylstra G, Kim E, Goyal AK (1997) Comparative molecular analysis of genes for polycyclic aromatic hydrocarbon degradation. Genet Eng 19: 257–269.

10 Microbial Degradation of Aliphatic and Aromatic Hydrocarbons with (Per) Chlorate as Electron Acceptor F. Mehboob1 . S. Weelink1 . F. T. Saia1 . H. Junca2 . A. J. M. Stams1 . G. Schraa1,* 1 Wageningen University and Research Center, Laboratory of Microbiology, Dreijenplein, Wageningen, The Netherlands *[email protected]. 2 AG Biodegradation Helmholtz-Zentrum fu¨r Infektionsforschung, Braunschweig, Germany 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 936 2 (Per)Chlorate Reduction and (Per)Chlorate-Reducing Microorganisms . . . . . . . . . . . . 937 3 Degradation of Aromatic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 937 4 Degradation of Aliphatic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 938 5 Genomics of (Per)Chlorate Reducing Bacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 941 6 Application in Soil Remediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 942 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 942

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_66, # Springer-Verlag Berlin Heidelberg, 2010

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Microbial Degradation of Aliphatic and Aromatic Hydrocarbons

Abstract: Aliphatic and aromatic hydrocarbons are rather persistent in anoxic conditions. These compounds are better degraded by aerobic microorganisms that use molecular oxygen for the initial activation. Addition of oxygen in a highly soluble form can help to remediate the soils polluted with such hydrocarbons. Perchlorate and chlorate are highly water soluble compounds that yields molecular oxygen upon microbial reduction. The oxygen produced may then be used to degrade persistent aliphatic and aromatic hydrocarbons. The metabolites thus formed are easier to degrade further anaerobically. Bacteria are known which can oxidize anaerobically recalcitrant hydrocarbons like benzene and alkanes coupled to chlorate reduction, but the present information is still scarce. By enrichment and isolation of (per) chlorate-reducing bacteria a better overview of the ability of these bacteria to degrade hydrocarbons can be obtained.

1

Introduction

Petroleum hydrocarbons belong to the most common environmental contaminants (Chayabutra and Ju, 2000). Major sources of petroleum hydrocarbon pollution are natural seepage and accidental spillage during transportation (www.eia.doe.gov). Petroleum hydrocarbons consist of four major components: saturates, aromatics, resins and asphaltenes (Head et al., 2006). On average, saturated and aromatic hydrocarbons together make 80% of the crude oil (Widdel and Rabus, 2001). The microbial degradation of hydrocarbons has been studied and reviewed extensively (Berthe-Corti and Fetzner, 2002; Heider, 2007; Leahy and Colwell, 1990; Wentzel et al., 2007; Widdel and Rabus, 2001). The mechanism of the aerobic microbial degradation of hydrocarbons is well described in several bacterial strains. During aerobic degradation, molecular oxygen not only acts as terminal electron acceptor but also as co-substrate (Berthe-Corti and Fetzner, 2002; Chayabutra and Ju, 2000; Leahy and Colwell, 1990). Oxygenases incorporate hydroxyl groups, derived from molecular oxygen, into the aliphatic chain or the aromatic ring. Aliphatics are usually converted to alcohols, while aromatics are generally metabolized to phenolic compounds (Heider et al., 1999), which are then mineralized to CO2 and water through common pathways like the TCA cycle. Insight in the anaerobic degradation of hydrocarbons is relatively new. The exact anaerobic mechanisms involved in the first steps of the activation of hydrocarbons are only described for few substrates and strains, like the mechanism of toluene degradation by Thauera aromatica K172, where the activation of toluene occurs via fumarate addition (Biegert et al., 1996). It appears that we are just starting to grasp the enormous diversity of mechanisms found in nature for the initial activation of hydrocarbons in anoxic environments. The range of substrates that can be utilized by the few microorganisms described with hydrocarbon degradation potential is narrow and the degradation rates are much lower than the ones of aerobic microorganisms (Chayabutra and Ju, 2000; Wentzel et al., 2007) as it would be assumed by evident thermodynamic constraints. Aerobic degradation seems to be the fastest option for mineralization of non-chlorinated hydrocarbons in the case of remediation strategies. Since anoxic zones may develop quickly after hydrocarbons pollution due to the activity of aerobic bacteria (Logan and Wu, 2002), the contaminated zone has to be removed physically/mechanically and treated elsewhere, or alternatively, oxygen has to be injected under high pressure to stimulate aerobic hydrocarbon degrading bacteria. The introduction of oxygen in the form of H2O2 is a widely used

Microbial Degradation of Aliphatic and Aromatic Hydrocarbons

10

technique, but hydrogen peroxide is toxic to many organisms and may instead inhibit the degradation of hydrocarbons (Morgen et al., 1993). This problem will not exist if oxygen can be introduced in an anoxic zone in a highly soluble alternative form.

2

(Per)Chlorate Reduction and (Per)Chlorate-Reducing Microorganisms

(Per)chlorate has a higher reduction potential than nitrate and oxygen (Coates and Achenbach, 2004; Stams et al., 2004) and has been suggested as an alternative electron acceptor in the oxidation of hydrocarbons (Coates et al., 1998). It also yields molecular oxygen upon reduction (Coates et al., 1999a; Rikken et al., 1996; Wolterink et al., 2002). The microbial reduction of perchlorate proceeds as follows:


ClO
4 ! ClO3 ! ClO2 ! Cl ! O 2

Perchlorate (ClO4
) is reduced to chlorate (ClO3
) through perchlorate reductase which in turn is reduced to chlorite (ClO2
) by chlorate reductase. Chlorite is then split into Cl
and O2 by chlorite dismutase (Rikken et al., 1996; Wolterink et al., 2002). This oxygen formation during anaerobic respiration is unique for (per)chlorate reducing bacteria (Coates et al., 1999a). Enzymes involved in this chlorate reduction pathway have been isolated and characterized (Kengen et al., 1999; Stenklo et al., 2001; van Ginkel et al., 1996). (Per)chlorate is considered to be anthropogenic, but a natural source of perchlorate are mineral deposits in Chile (Orris et al., 2003). Perchlorate is used as rocket fuel, while chlorate is used for making matches and as herbicide (Gullick et al., 2001; Urbansky, 1998). Since perchlorate salts are highly soluble in water (sodium perchlorate solubility is 2 kg/l), they are readily transported through surface water and groundwater (Xu et al., 2003). Growth rates and cell yields on (per)chlorate are comparable with aerobic ones and no toxic intermediates accumulate (Logan, 2001). Microbial (per)chlorate reduction has been reviewed by Coates and Achenbach (2004), Logan and Wu (2001), Xu et al. (2003). Perchlorate reducing bacteria are ubiquitous and phylogenetically diverse (Coates et al., 1999b) and have been isolated from pristine as well as contaminated soils and sediments (Balk et al., 2008; Kesterson et al., 2005; Romanenko et al., 1976; Stepanyuk et al., 1992; Wolterink et al., 2002, 2005). Culture-dependent evidence for the ubiquity of (per)chlorate reducing bacteria came from most probable number count assays in samples from a variety of soils and sediments, where the numbers of (per)chlorate reducing bacteria ranged from 2  103 to 2  106 cells per gram (Coates et al., 1999b). Kesterson et al. (2005) showed that (per)chlorate reducing bacteria could even be two orders of magnitude higher in some locations. The majority of perchlorate reducing bacteria are facultative anaerobic mesophiles and have a pH optimum close to neutral.

3

Degradation of Aromatic Compounds

Aromatic hydrocarbons like BTEX are of great concern as contaminants in soil, due to their solubility, mobility, toxicity and persistence under anaerobic conditions (Coates et al., 1999a). Only few bacterial strains have been isolated that are able to grow on BTEX anaerobically (Rabus and Widdel 1995; Kniemeyer et al., 2003) except for toluene. None of these isolates is able to degrade all the BTEX compounds (Chakraborty et al., 2005). Two denitrifying, benzene

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10

Microbial Degradation of Aliphatic and Aromatic Hydrocarbons

degrading bacteria, Dechloromonas aromatica strain RCB and strain JJ have recently been isolated. Strain RCB was enriched on 4-chlorobenzoate and uses chlorate as an electron acceptor (Coates et al., 2001). In addition to benzene, this strain was able to grow on toluene, all the isomers of xylene and ethylbenzene. The rates of benzene degradation were slightly higher when oxygen or (per)chlorate were used as electron acceptor instead of nitrate (Chakraborty et al., 2005). Two denitrifying bacteria, Azoarcus sp. strain DN11 and strain AN9, were reported to grow on benzene as sole source of carbon and energy (Kasai et al., 2006; 2007), but these strains were not tested on (per)chlorate. We isolated a bacterium Alicycliphilus strain BC from a chlorate-reducing community, that had 20–1,650 times higher benzene degradation rates than reported for anaerobic benzene degradation (Weelink et al., 2007). This bacterium is able to grow on benzene, toluene, phenol, and catechol with chlorate and oxygen, but not with nitrate (Weelink et al., 2008). This suggests that during growth on chlorate, the oxygen that is formed via chlorite dismutation is incorporated by an oxygenase in the aromatic ring. Some characteristics of pure cultures that degrade benzene anaerobically are presented in > Tables 1 and 2. Another likely mechanism in the enrichment obtained by Weelink et al. (2007) is interspecies oxygen transfer. A bacterium related to Mesorhizobium sp. WG dominates the enrichment when grown on benzene and oxygen. Another bacterium, related to Stenotrophomonas acidaminophila, is enriched when the enrichment culture is supplied with acetate and chlorate. It seems that the latter one is a chlorate reducer, which forms oxygen during chlorite dismutation, and the former one is an aerobic benzene degrader, which utilizes this oxygen to degrade benzene (Weelink et al., 2007). In another study, 14C labeled naphthalene was quickly converted under anoxic conditions to 14CO2. Chlorite and not (per)chlorate was directly added to a washed whole cell suspension of a (per)chlorate reducing, non-hydrocarbon degrading bacterium, D. agitatus strain CKB, and the hydrocarbon oxidizing Pseudomonas strain JS150 (Coates et al., 1999a). They even found higher degradation rates under anoxic conditions compared to aerobic controls which was attributed to the limited diffusion rate of oxygen. In a study by Logan and Wu (2002), toluene degradation rates were 1.36 fold enhanced when chlorate was amended to a sand column. This increase in toluene degradation was attributed to oxygen formation by chlorate-reducing organisms which is taken up by toluene degrading bacteria to hydroxylate the toluene ring through oxygenases. In another soil column study, high benzene degradation rates (31 mmol/l/hr) coupled to chlorate reduction were observed. The results with batch cultures showed that only chlorate and not nitrate was used as an electron acceptor. This again points to an involvement of an oxygenase mediated mechanism of action (Tan et al., 2006).

4

Degradation of Aliphatic Compounds

Alkanes are chemically not very reactive due to lack of functional groups, the presence of only sigma bonds, the non-polar nature and the very low solubility in water. Although n-alkanes are degraded faster than the other components of the petroleum hydrocarbons, they are quantitatively the major fraction of crude oil (Head et al., 2006). The exact mechanisms involved in the first steps of the activation of alkanes in anoxic conditions are not yet completely understood. Recently, a mechanism similar to toluene activation through fumarate addition has been proposed for the degradation of alkanes and genes identical to toluene degradation have been found (Grundmann et al., 2008). A different mechanism of initial

Wastewater treatment plant

Contaminated groundwater

Contaminated groundwater

Alicycliphilus strain BC

Azoarcus strain DN 11

Azoarcus strain AN 9

Benzene + O2 on dCGY medium

Benzene + NO3
on dCGYb medium

b

ND

ND ND

ND

1–2  0.6 30–37

Rod

Benzene + ClO3
with 0.125 g/L FYE

25

30

30

1.8  0.5

Rod AHDSa + NO3
(acetate as C source)

2,6-anthrahydroquinone disulphonate dCGY medium (Bact. Casamino acid, Glycerol, Yeast extract, Agar) c Estimated from > Fig. 2 (Chakraborty et al., 2005) d Estimated from > Fig. 4 (Kasai et al., 2006)

a

Lake Sediments

Dechloromonas strain JJ

30

1.8  0.5

Rod

River sediment 4 – Chlorobenzoate + ClO3


Dechloromonas aromatica strain RCB

Isolated on

Isolated from

Name of Strain

Opt. Temp. Shape Size (um) ( C)

ND

7

7.3

7.2

7.2

Opt. pH

ND

15 13 daysd (benzene + NO3
)

9 daysd 15 (benzene + NO3
)

1.4 days 1,000 (benzene + ClO3
)

ND

160 4 days (benzene + ClO3
)

c

Doubling time

Highest conc. of benzene tested. (uM)

Kasai et al. (2006)

Kasai et al., 2006; 2007

Weelink et al., (2008)

Coates et al. (2001)

Coates et al., 2001; Chakraborty et al., 2005

References

. Table 1 Comparison of selected features reported for all the isolates (all belong to subclass b-proteobacteria) described to be capable of anaerobic benzene degradation

Microbial Degradation of Aliphatic and Aromatic Hydrocarbons

10 939

+

+

+

nd

nd

nd

Acetate

Toluene

Benzene

Phenol

Catechol

Benzoate

+

nd

+

+

+

+

NO3


nd

nd

nd

+

+

+

ClO3


+

nd

nd

+

+

+

O2

+

nd

nd

+

+

+

NO3


nd

nd

nd

nd

nd

nd

ClO3


Dechloromonas strain JJ










+

+

+

+





+




+

+

ClO3





NO3


+

+

+

+

O2

Alicycliphilus strain BC

a 20%–90% of amended substrate degraded nd not described/not detected; + growth;
no growth (References for organisms can be found in > Table 1)

O2

Substrates

Dechloromonas aromatica RCB

+

nd




+

nd

nd

O2

+

nd




+a

+

nd

NO3


nd

nd

nd

nd

nd

nd

ClO3


Azoarcus strain DN 11

nd

nd

nd

+a

nd

nd

O2

nd

nd

nd

+

+

nd

NO3


nd

nd

nd

nd

nd

nd

ClO3


Azoarcus strain AN 9

10

. Table 2 Comparison of degradation of hydrocarbons under different electron accepting conditions reported for isolates described to be capable of anaerobic benzene degradation

940 Microbial Degradation of Aliphatic and Aromatic Hydrocarbons

Microbial Degradation of Aliphatic and Aromatic Hydrocarbons

10

. Figure 1 Proposed mechanism of degradation of alkanes coupled to chlorate reduction. Oxygen released from chlorite dismutation is used by the oxygenase enzyme to incorporate it inside the alkane molecule to form an alcohol.

carboxylation has been proposed for the alkane degrading sulfate reducing strain Hxd 3 (Aeckersberg et al., 1998; Callaghan et al., 2009). Alkane degradation studies under oxygen limited conditions show that oxygen is required only for initial activation of alkanes and further degradation can occur anaerobically (Chayabutra and Ju, 2000; Michaelsen et al., 1992). So it might be possible that (per)chlorate reducing bacteria having an alkane oxygenase enzyme could utilize the oxygen produced during the chlorite dismutation step and incorporate it in the alkane to activate it and form an alcohol. The alcohol is then further degraded by beta oxidation. Recently, we found that Pseudomonas chloritidismutans AW-1T, previously isolated on acetate and chlorate (Wolterink et al., 2002), is able to grow with medium-length n-alkanes with chlorate as electron acceptor. This strain degrades alkanes with oxygen and chlorate, but not with nitrate, suggesting the involvement of oxygenases in the breakdown of alkanes with chlorate (Mehboob et al., 2009). The theoretical proposed reaction mechanism is presented in > Fig. 1. The degradation of alkanes may also be tentatively possible through interspecies oxygen transfer, i.e. part of the oxygen produced by a (per)chlorate reducer through chlorite dismutation may be used by an alkane degrader for incorporation inside the alkane.

5

Genomics of (Per)Chlorate Reducing Bacteria

The genetics of (per)chlorate reduction has not yet been studied in detail. Transcriptional analysis of the chlorite dismutase gene (cld ) in D. agitata revealed that the cld gene was transcriptionally up-regulated when grown anaerobically on (per)chlorate (Bender et al., 2002). In another study, the possible horizontal transfer of chlorite dismutase (cld) genes, based upon their gene phylogeny and the host taxonomy affiliation was suggested(Bender et al., 2004).

941

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Thorell et al. (2003) demonstrated that chlorate reductase and chlorite dismutase are clustered together in close proximity in Ideonella dechloratans suggesting an evolution towards a functional succession. One known (per)chlorate reducer, D. aromatica strain RCB, has been fully sequenced (http://genome.jgi-psf.org/finished_microbes/decar/decar.home.html). Chlorate reductases in most of the genomes are annotated as nitrate reductases as both enzymes belong to same dimethyl sulfoxide (DMSO) reductase family and can reduce each others substrate. The chlorite dismutase gene in the genome of strain RCB is clustered together with the nitrate reductase showing their functional connection. Strain RCB contains a variety of annotated oxygenases which may metabolize hydrocarbons, such as toluene-4-mono-oxygenase system B, 2-nitropropane dioxygenase and subunits of extradiol ring cleavage dioxygenases, aromatic ring hydroxylating dioxygenases, alpha subunits of Rieske (2Fe-2S) ring hydroxylating oxygenases, FAD dependent monooxygenases, phenyl acetate-CoA oxygenases and many other putative oxygenases (http://www.ncbi.nlm.nih.gov). Similarly, Alicycliphilus strain BC has a putative benzene mono-oxygenase (BC-BMOa) gene and a putative catechol 2,3- dioxygenase (BC-C23O) gene. The BC-BMOa gene sequence of A. denitrificans strain BC is 76% identical to the D. aromatica strain RCB sequence. Although these sequences are putative, all sequence features and physiological data support the statement that these sequences are active and encode functional proteins (Weelink et al., 2008). The frequent distribution of chlorite dismutase genes (Maixner et al., 2008) and presence of many environmentally relevant oxygenases inside the genome of (per)chlorate-reducing bacteria indicate the unexploited power and the unexplored potential of chlorate reducers for bioremediation processes perspectives.

6

Application in Soil Remediation

To our knowledge, only one study has been performed for the application of (per)chlorate reduction in soil remediation. 14C labeled benzene was added in this study to anoxic soil samples that were pre-treated with Dechloromonas agitatus strain CKB, a non hydrocarbon degrading (per)chlorate reducer, and chlorite dismutating strain. The indigenous hydrocarbon degrading population was rapidly stimulated and more than 90% of 14C labeled benzene was recovered as 14CO2. The rates were similar to aerobic controls (Coates et al., 1999a). Bioremediation of soils contaminated with hydrocarbons depend on many factors including availability of the hydrocarbons, presence of nutrients and electron acceptors, temperature, salinity, water activity and pH (Leahy and Colwell, 1990). Since most of the (per)chlorate reducing organisms have a pH optimum close to neutral, the pH may have to be adjusted before the actual bioremediation starts. The addition of fertilizers along with the soluble electron acceptor ((per)chlorate) addition may enhance the remediation process in soils lacking nutrients. The addition of (per)chlorate alone may not be sufficient. Therefore a dosage of a readily degradable substrate like acetate to enhance (per)chlorate reduction and production of oxygen may be essential (Tan et al., 2006).

7

Research Needs

A solution for the biological remediation of hydrocarbons contaminated soils where oxygen is the main limiting factor is the introduction of oxygen in a highly soluble form. (Per)chlorate, which acts both as electron acceptor and as oxygen source, is an appealing compound. It has a

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high solubility, a redox potential comparable to oxygen, and there are numerous (per)chlorate reducing bacteria in nature. We are at the beginning of deciphering and understanding the diversity, relevance and application of per(chlorate)-reducing bacteria that can degrade hydrocarbons and other compounds, which are rather recalcitrant under anoxic conditions. Thus, this aspect in the study of the ecology of biodegradation deserves continued investigation efforts. Only a few hydrocarbon-degrading chlorate reducing bacteria have been enriched and isolated up to now. Considering the enormous potential of (per)chlorate reducing bacteria, there is a need to enrich and isolate (per)chlorate-reducing bacteria over a broader range of hydrocarbons. In addition, most of the known (per)chlorate reducers have been isolated on simple substrates like fatty acids so further screening of these bacteria for hydrocarbon degradation is suggested. Similarly metagenome analysis may shed light on the occurrence and distribution of genes linked to the oxygenase-dependent breakdown of recalcitrant compounds and to (per)chlorate reduction. This may also provide insight into the biodegradation potential of polluted soils.

Acknowledgments We would like to thank Dutch Center for Soil Quality Management and Knowledge Transfer, (SKB; www.skbodem.nl) The Netherlands, Wageningen Institute for Environment and Climate Research (WIMEK), The Netherlands, Fundac¸a˜ode Amparo a` Pesquisa do Estado de Sa˜o Paulo (FAPESP) Brazil and Higher Education Commission of Pakistan (HEC; www.hec.gov.pk/) for providing funds for the present study.

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11 Hydrocarbon Degradation Coupled to Metal Reduction M. L. Heinnickel . F. M. Kaser . J. D. Coates* Department of Plant and Microbial Biology, University of California, Berkeley, CA, USA *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 948 2 Hydrocarbon Degradation by G. metallireducens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 948 3 Conversion of Benzoyl-CoA to Cyclohexa-1,5-Dienecarboxy-CoA . . . . . . . . . . . . . . . . . . 949 4 A Unique Benzoyl-CoA Reductase in Geobacter . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 950 5 Activation of Benzoyl-CoA in G. metallireducens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 952 6 Catabolism of Cyclohexa-1,5-Dienecarboxy-CoA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 953 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 953

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_67, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: It is known that hydrocarbon degradation can be coupled to dissimilatory microbial Fe(III) reduction under anaerobic conditions. Of the phylogenetically diverse Fe(III)reducing organisms isolated and characterized only the Geobacter species have been shown to be capable of hydrocarbon oxidation, and even then, these organisms could only utilize simple monoaromatic hydrocarbons such as toluene. Analysis of the Geobacter metallireducens genome suggests that the route of aromatic hydrocarbon degradation in Fe(III) reducing bacteria mimics that found in facultative anaerobes with a few exceptions. Similar to facultative anaerobes, parent aromatic hydrocarbons are converted through an upper pathway to benzoyl-CoA. However, benzoyl-CoA appears to be reduced to cyclohexa-1,5-dienecarboxyCoA by a novel protein complex. This novel benzoyl-CoA reductase (BCR) is hypothesized to contain proteins that have similarities to those found in hydrogenases, heterodisulfide reductases, NADH: Ubiquinone oxidoreductases, aldehyde/ketone: ferredoxin oxidoreductases, and selenium-containing proteins. Unlike other organisms, the method of activation is not ATP dependant, and may occur by a novel process. Here we discuss possible methods of activation, along with a mechanism detailing the steps of benzoate dearomatization and differences in benzoyl-CoA reductases from G. metallireducens and Thauera aromatica, a well-characterized hydrocarbon-oxidizing facultative anaerobe.

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Introduction

Several previous studies have demonstrated the hydrocarbon biodegradative capacity of dissimilatory Fe(III) reducing bacteria (Lovley and Lonergan, 1990; Lovley et al., 1994, 1996; Anderson et al., 1998; Anderson and Lovley, 1999; Rooney-Varga et al., 1999; Coates et al., 2001; Kunapuli et al., 2007) and various aromatic hydrocarbons have been shown to be biodegraded in sediments in which Fe(III) was the terminal electron acceptor (Lovley et al., 1994, 1996; Anderson et al., 1998; Anderson and Lovley, 1999; Kunapuli et al., 2007). Microbial community studies demonstrated that these sediments were often enriched in organisms of the family Geobacteraceae (Rooney-Varga et al., 1999). However, there was no direct evidence to show that members of this family of organisms are capable of hydrocarbon metabolism other than toluene degradation. In contrast, stable isotope analysis of a benzenedegrading Fe(III)-reducing enrichment culture obtained from soil of a former coal gasification site in Gliwice, Poland, suggested that hydrocarbon degradation was mediated by a novel syntrophic interaction between members of the gram-positive clostridial Peptococcaceae and the Desulfobulbaceae species within the delta subclass of the Proteobacteria (Kunapuli, Lueders et al., 2007). Of all the known phylogenetically diverse Fe(III)-reducing bacteria only Geobacter metallireducens and G. grbiciae are capable of hydrocarbon degradation in pure culture and even then their metabolism is limited to toluene (Lovley and Lonergan, 1990; Coates et al., 2001).

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Hydrocarbon Degradation by G. metallireducens

The degradation of aromatic compounds including toluene by G. metallireducens can be inferred through its available genome sequence. Computational annotation indicates that the organism has genes that enable the degradation of phenol, p-cresol, benzyl alcohol, benzaldehyde, toluene, and 4-hydroxybenzoate (Butler et al., 2007). Similar to facultative

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anaerobes, it is believed that benzoyl-CoA is a common intermediate in the degradation of all listed aromatic compounds under Fe(III)-reducing conditions. In addition, many of the genes involved in the processing of these compounds are found on the ‘‘aromatic island’’; a region of genes in the G. metallireducens genome that are believed to be involved in aromatic hydrocarbon degradation (Butler et al., 2007). Biochemical analysis on the enzyme responsible for the conversion of toluene to benzylsuccinate, an activating step necessary for toluene conversion to benzoyl-CoA, revealed the initial activation is mediated by a glycine-radical-dependent enzyme called benzyl succinate synthase (BSS) (Leuthner et al., 1998). A similar type reaction is observed in toluenedegrading denitrifying bacteria, sulfate reducing bacteria, phototrophic bacteria, and methanogenic consortia (Chakraborty and Coates, 2003). As in T. aromatica (Leuthner et al., 1998), BSS was identified in G. metallireducens by its activity, the radical catalyzed stereospecific addition of fumurate to toluene to produce (R)-benzylsuccinate (Kane et al., 2002). In addition, bssA and bssB, genes for the subunits of this enzyme complex, were identified using southern blot hybridization (Kane et al., 2002). The C-terminal region of BssA from Thauera aromatica str. K172 corresponded strongly to 411 residues found in the BssA from G. metallireducens (Kane et al., 2002). Most notably, this subunit was found to contain a conserved cysteine and glycine. The cysteine is believed to participate in the formation of a radical on the amino acid glycine (Kane et al., 2002). This essential glycine-radical abstracts a hydrogen atom from toluene forming a benzyl radical. This enzyme bound benzyl radical adds to the double bond of the co-substrate fumarate, which is also bound at the active site. The resulting benzylsuccinyl radical abstracts the hydrogen from the aforementioned glycine residue, thus regenerating the glycine radical that initiated the reaction (Kane et al., 2002). The BSS from G. metallireducens appears to follow this model reaction as d8-toluene (toluene deuterated at all positions) was converted to d8-benzylsuccinate, suggesting the initial deuterium atom removed from toluene to form the benzyl radical was subsequently added back to the benzylsuccinyl radical to form benzylsuccinate (Kane et al., 2002).

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Conversion of Benzoyl-CoA to Cyclohexa1,5-Dienecarboxy-CoA

As outlined above, a central metabolite of aromatic degradation in Fe(III) reducing bacteria is benzoyl-CoA. Benzoyl-CoA is formed by benzoyl-CoA ligase (Egland et al., 1995; Schuhle et al., 2003). This enzyme hydrolyzes ATP to AMP and PPi while ligating benzoate to coenzyme A through a thioester linkage (Egland et al., 1995). While an energetically intensive step (it uses two equivalents of ATP per unit benzoate), it is consistently seen in all strictly anaerobic bacteria that oxidize aromatics, as well as facultative anaerobes (Elder and Kelly, 1994; Fuchs, 2008). Analysis of the upregulated proteome of G. metallireducens in the presence of benzoate showed that one such upregulated protein, called BamY, had greater than 50% sequence identity to benzoyl-CoA ligases from T. aromatica, Azoarcus evansii, and Magentospirillum magnetotacticum (Wischgoll et al., 2005). The gene for this protein was cloned and overexpressed in Escherichia coli (Wischgoll et al., 2005). Using a spectrophotometric assay for carboxylic acid coenzyme A ligases, overexpressed BamY was found to have a specific activity of 16 mmol min 1 mg 1 for benzoate. This value was higher than that reported for other anaerobes, and proved the existence of this enzyme in Fe(III) reducing organisms (Wischgoll et al., 2005).

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The next step in the aromatic degradation pathway appears to be a two electron reduction of the aromatic ring. In T. aromatica this step is performed by the enzyme benzoyl-CoA reductase (BCR) (Boll and Fuchs, 1995; Boll et al., 1997). BCR is composed of four subunits (bcrA-D) that contain 3 [4Fe-4S] clusters and two ATP-binding domains (Boll et al., 1997; Breese et al., 1998; Boll et al., 2000; Boll et al., 2001) (also see > Fig. 1). Similar to nitrogenase, BCR couples electron transfer to its substrate with ATP hydrolysis (Boll et al., 1997; Mobitz et al., 2004). In this reaction ATP binds to BCR changing the enzyme from a ‘‘closed’’ to an ‘‘open’’ state (Mobitz et al., 2004). In the ‘‘open’’ state, a unique [4Fe-4S] cluster is reduced by soluble ferredoxin, and benzoyl-CoA enters the enzyme’s substrate-binding pocket (Mobitz et al., 2004). Following these two events, the ATP bound to the enzyme is hydrolyzed. This ATP hydrolysis serves as an ‘‘activating’’ step changing the spin state of the reduced [4Fe-4S] cluster from S = 1/2 to S = 7/2 (Boll et al., 2000). In addition, this hydrolysis also presumably lowers the redox potential of this cluster to the 1 V redox potential required for the reduction of benzoyl-CoA to a dienoyl-CoA radical anion (Boll, 2005; Fuchs, 2008). It appears the two electron reduction of benzoyl-CoA occurs in G. metallireducens (Peters et al., 2007). A study by Wischoll et al. showed that 44 genes upregulated during benzoate degradation in G. metallireducens included a putative cyclohexa-1,5-dienecarboxy-CoA dehydratase (bamR). This enzyme hydrolyzes the product of benzoyl-CoA reductase, cyclohexa1,5-dienecarboxy-CoA, indicating that the initial two electron reduction occurs (Peters et al., 2007). In addition, whole cell lysate from G. metallireducens was shown to have the ability to oxidize cyclohexa-1,5-dienecarboxy-CoA, but not cyclohexa-1-ene-carbonyl-CoA (a metabolite formed when phototrophs oxidize benzoyl-CoA by a presumably different pathway (Harwood et al., 1999)). Further, when the bamR gene product is overexpressed in E. coli, it was capable of oxidizing cyclohexa-1,5-dienecarboxy-CoA to 6-OH-cyclohexanoyl-CoA (Peters et al., 2007). This reaction is characteristic of the pathway exemplified by T. aromatica.

4

A Unique Benzoyl-CoA Reductase in Geobacter

Unexpectedly, no transcript bearing any similarity to benzoyl-CoA reductase is upregulated during growth on benzoate (Butler et al., 2007). Although one gene in the G. metallireducans genome bears some significant similarity to one of the subunits in benzoyl-CoA reductase (Hosoda et al., 2005), it was not upregulated during growth on benzoate (Butler et al., 2007). Furthermore, G. metallireducens whole cell lysate cannot reduce benzoyl-CoA in the presence of ATP and added reductants (Wischgoll et al., 2005). Therefore it appears that G. metallireducens reduces benzoyl-CoA to cyclohexa-1,5-dienecarboxy-CoA with a novel enzyme. This alternative pathway may exist as a method to save ATP (Fuchs, 2008) as obligate anaerobes only acquire a maximum of four ATPs per molecule of benzoate oxidized (Fuchs, 2008). This, in combination with the fact that G. metallireducens hydrolyzes two molecules of ATP to ligate benzoate to coenzyme A (Wischgoll et al., 2005), suggests that there is a limited net energy gain from benzoate metabolism. Therefore, it may make sense for the organism to utilize an alternative benzoyl-CoA reductase that does not expend ATP. Analysis of the 44 upregulated genes in G. metallireducens in the presence of benzoate indicated that many previously uncharacterized enzymes were expressed (Wischgoll et al., 2005). Some of these enzymes appeared to contain selenium and tungsten or molybdenum (Wischgoll et al., 2005). The presence of selenium in an enzyme essential to this process has previously been indicated with other strictly anaerobic bacteria oxidizing hydrocarbons (Peters et al., 2004;

. Figure 1 A diagram showing the steps of dearomatization of benzoate in Geobacter metallireducens. The step catalyzed by a different enzyme complex compared to T. aromatica is indicated. The numbers 1–6 refer to intermediates in the dearomatization of benzoate. They are 1 – benzoate; 2 – benzoyl-CoA; 3 – cyclohexa-1,5-diene-1-carbonyl-CoA; 4 – 6-hydroxylcyclohex-1-en-1-carbonyl-CoA dehydrogenase; 5 – 6-oxocyclohex-1-ene-1-carbonyl-CoA; 6 – 3hydroxypimelyl-CoA. The letters A–D refer to enzymes involved in the benzoate dearomatization process. They are A – benzoyl-CoA ligase; B – cyclohexa1,5-diene-1-carbonyl-CoA dehydratase; C – 6-hydroxylcyclohex-1-en-1-carbonyl-CoA; and D – 6-oxocyclohex-1-ene-1-carbonyl-CoA hydratase.

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Wischgoll et al., 2005). In 2004, it was found that D. multivorans, a sulfate reducing bacteria whose hydrocarbon degradation pathway may bear similarity to the pathway seen in G. metallireducens, has an absolute requirement for molybdenum and selenium to degrade benzoate (Peters et al., 2004). It was also found that two proteins with molecular weights of 30 and 100 kDa were upregulated in D. multivorans in the presence of benzoate and could be labeled with 75Se (Peters et al., 2004). This selenium remained even after denaturing the proteins with SDS, prior to analysis on a polyacrylamide gel. The tenacity of this label suggests that the selenium is most likely incorporated into these polypeptides as selenocysteine. One of the polypeptides upregulated when G. metallireducens is incubated with benzoate, BamF, is similar to VhuD (Wischgoll et al., 2005; Butler et al., 2007), a protein in methanogens that binds selenocysteine. VhuD has been implicated in the formation of a tight complex between soluble hydrogenases and heterodisulfide reductases (Stojanowic et al., 2003). Additional polypeptides produced in the presence of benzoate have similarities to subunits of heterodisulfide reductases, soluble hydrogenases, and NADH:Ubiquinone oxidoreductases (BamC-I) (Wischgoll et al., 2005). The protein BamC has a strong similarity to soluble hydrogenases that contain three [4Fe-4S] clusters, but the whole cell lysate of G. metallireducens does not have any hydrogenase properties. In addition, no large subunit containing the NiFe center required for proton reduction is present in the G. metallireducens genome (Wischgoll et al., 2005). The proteins BamD-E contain Fe/S clusters and flavins, and are similar to heterodisulfide redutases (Wischgoll et al., 2005). It could be that BamC-E form a tight complex that transfers electrons to a potential catalytic subunit (> Fig. 1). This tight complex would be anchored by the BamF protein, functioning similarly to VhuD (Wischgoll et al., 2005). The catalytic subunit in this case may be formed by BamB. The gene for BamB is upregulated 36-fold in the presence of benzoate (Butler et al., 2007). This gene is similar to molybdenum containing aldehyde:ferredoxin oxidoreductases, and formaldehyde:ferredoxin oxidoreductases (Wischgoll et al., 2005; Butler et al., 2007). These enzymes typically contain a bismolybdo(tungsto)pterin mononucleotide cofactor and a [4Fe-4S] cluster (Chan et al., 1995; McMaster and Enemark, 1998). The motifs for these cofactors are present in BamB (Wischgoll et al., 2005), implying that they may be involved in the reduction of benzoyl-CoA (see > Fig. 1). This implication appears to fit with present data that molybdenum is a required cofactor for the oxidation of benzoate in G. metallireducens (Wischgoll et al., 2005). In addition, molybdenum biosynthetic genes are upregulated in the presence of benzoate in G. metallireducens (Butler et al., 2007). Other proteins expressed in the presence of benzoate include BamG-I, which bear similarity to NAD(P)H:Ubiquinone oxidoreductases (Wischgoll et al., 2005; Butler et al., 2007). It is entirely possible that the BamG-I complex oxidizes NAD(P)H and transfers these electrons to the BamB subunit through the BamC-F complex (see > Fig. 1). These electrons would be used to reduce benzoyl-CoA to cyclohexa-1,5dienecarboxy-CoA.

5

Activation of Benzoyl-CoA in G. metallireducens

The challenging question with this hypothesis remains the method of activation. NAD(P)H has redox potential of 340 mV, while the redox potential of the dienoyl CoA radical anion that results from a single reduction of benzoyl-CoA has a redox potential of approximately 1 V. This disparity in redox potential of over 600 mV can be accomplished by coupling the electron transfer to an activation step. In the benzoyl-CoA reductase from T. aromatica this

Hydrocarbon Degradation Coupled to Metal Reduction

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activation step occurs by the hydrolysis of ATP. However, experiments with whole cell lysates of G. metallireducens indicate that ATP and reductants alone cannot stimulate the reduction of benzoyl-CoA (Wischgoll et al., 2005). Further none of the aforementioned polypeptides have ATP-binding domains (Wischgoll et al., 2005), indicating a different method of activation may take place. One possible solution is that this reduction of benzoyl-CoA is driven by some membrane potential (Wischgoll et al., 2005). In addition, the benzoyl-CoA reductase from G. metallireducens may have an alternative mechanism of stabilizing the dienoyl-CoA radical that forms following an electron transfer event. This different mechanism may decrease the energy required to activate the enzyme prior to this reduction.

6

Catabolism of Cyclohexa-1,5-Dienecarboxy-CoA

Following the reduction of benzoyl-CoA in T. aromatica, the resulting cyclohexa-1,5dienecarboxy-CoA is converted to 3-hydroxypimelyl-CoA. The enzymes required for this conversion; cyclohexa-1,5-diene-1-carbonyl-CoA dehydratase, 6-hydroxylcyclohex-1-en-1carbonyl-CoA dehydrogenase, and 6-oxocyclohex-1-ene-1-carbonyl-CoA hydratase all have homologues in the G. metallireducens genome (Butler et al., 2007). All of these genes are upregulated in the presence of benzoate (Butler et al., 2007). Presumably, the resulting 3-hydroxypimelyl-CoA enters the b-oxidation pathway, where it is converted to acetyl-CoA subunits. Acetyl-CoA subunits are transformed into CO2 and NADH by the tricarboxylic acid cycle. A genomic analysis also shows that the enzymes for this cycle change slightly in the presence of benzoate. One additional adaption by G. metallireducens may generate more ATP per unit benzoate. Acetate: succinyl-CoA transferase, which functions in the G. sulfurreducens to transfer coenzyme A from succinyl-CoA to acetate, is downregulated in G. metallireducens 2.2-fold in the presence of benzoate (Butler et al., 2007). However, a homologue of succinyl-CoA synthetase was found to be upregulated and appears to convert succinyl-CoA to succinate, coupled to a substrate level phosphorylation (Butler et al., 2007). One of three copies of this gene is found in the ‘‘aromatic island’’ in the G. metallireducens sequence, and is upregulated 8- to 32-fold in the presence of benzoate. This upregulation of succinyl-CoA synthase appears prudent as acetyl-CoA is already formed from b-oxidation, and there is no need to acquire coenzyme A groups from succinyl-CoA. This is an interesting modification that generates additional molecules of ATP during the degradation of benzoate.

7

Research Needs

Key enzymes involved in processing benzene remain unknown, although it appears clear that benzene oxidation is coupled to Fe(III) reduction in the environment by some unknown organism. Isolating a pure culture or well-characterized consortia that can oxidize benzene coupled to Fe(III) reduction would be a large contribution to this field. Future directions for this field involve a complete characterization of genes involved in benzoyl-CoA reduction. The absence of homologues to the benzoyl-CoA reductase protein complex indicates that a novel enzyme system is involved in the reduction of benzoyl-CoA. The necessary requirement of both selenium and molybdenum in benzoate degradation implies this enzyme complex may contain these atoms. Proteins related to hydrogenases,

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heterodisulfide reductases, NADH:Ubiquinone oxidoreductases, Aldehyde/ketone:ferredoxin oxidoreductases, and a selenium-containing protein have all been implicated as possible components in this novel complex. Another key question regarding this enzyme complex concerns a method of activation of electrons entering the complex from physiological sources.

References Anderson RT, Lovley DR (1999) Naphthalene and benzene degradation under Fe(III)-reducing conditions in petroleum-contaminated aquifers. Biorem J 3(2): 121–135. Anderson RT, Rooney-Varga JN, et al. (1998) Anaerobic benzene oxidation in the Fe(III) reduction zone of petroleum contaminated aquifers. Environ Sci Technol 32(9): 1222–1229. Boll M (2005) Key enzymes in the anaerobic aromatic metabolism catalysing Birch-like reductions. Biochim Biophys Acta 1707(1): 34–50. Boll M, Fuchs G (1995) Benzoyl-coenzyme A reductase (dearomatizing), a key enzyme of anaerobic aromatic metabolism. ATP dependence of the reaction, purification and some properties of the enzyme from Thauera aromatica strain K172. Eur J Biochem 234(3): 921–933. Boll M, Albracht SS, et al. (1997) Benzoyl-CoA reductase (dearomatizing), a key enzyme of anaerobic aromatic metabolism. A study of adenosinetriphosphatase activity, ATP stoichiometry of the reaction and EPR properties of the enzyme. Eur J Biochem 244(3): 840–851. Boll M, Fuchs G, et al. (2000) EPR and Mossbauer studies of benzoyl-CoA reductase. J Biol Chem 275(41): 31857–31868. Boll M, Fuchs G, et al. (2001) Single turnover EPR studies of benzoyl-CoA reductase. Biochemistry 40(25): 7612–7620. Breese K, Boll M, et al. (1998) Genes coding for the benzoyl-CoA pathway of anaerobic aromatic metabolism in the bacterium Thauera aromatica. Eur J Biochem 256(1): 148–154. Butler JE, He Q, et al. (2007) Genomic and microarray analysis of aromatics degradation in Geobacter metallireducens and comparison to a Geobacter isolate from a contaminated field site. BMC Genomics 8: 180. Chakraborty R, Coates JD (2003) Anaerobic degradation of monoaromatic hydrocarbons. Appl Microbiol Biotechnol 64: 437–446. Chan MK, Mukund S, et al. (1995) Structure of a hyperthermophilic tungstopterin enzyme, aldehyde ferredoxin oxidoreductase. Science 267(5203): 1463–1469.

Coates JD, Bhupathiraju VK, et al. (2001) Geobacter hydrogenophilus, Geobacter chapellei and Geobacter grbiciae, three new, strictly anaerobic, dissimilatory Fe(III)-reducers. Int J Syst Evol Microbiol 51: 581–588. Egland PG, Gibson J, et al. (1995) Benzoate-coenzyme A ligase, encoded by badA, is one of three ligases able to catalyze benzoyl-coenzyme A formation during anaerobic growth of Rhodopseudomonas palustris on benzoate. J Bacteriol 177(22): 6545–6551. Elder DJ, Kelly DJ (1994) The bacterial degradation of benzoic acid and benzenoid compounds under anaerobic conditions: unifying trends and new perspectives. FEMS Microbiol Rev 13(4): 441–468. Fuchs G (2008) Anaerobic metabolism of aromatic compounds. Ann N Y Acad Sci 1125: 82–99. Harwood CS, Burchhardt, G, et al. (1999) Anaerobic metabolism of aromatic compounds via the benzoyl-CoA pathway. FEMS Microbiol Rev 22: 439–458. Hosoda A, Kasai Y, et al. (2005) Development of a PCR method for the detection and quantification of benzoyl-CoA reductase genes and its application to monitored natural attenuation. Biodegradation 16(6): 591–601. Kane SR, Beller HR, et al. (2002) Biochemical and genetic evidence of benzylsuccinate synthase in toluenedegrading, ferric iron-reducing Geobacter metallireducens. Biodegradation 13(2): 149–154. Kunapuli U, Lueders T, et al. (2007) The use of stable isotope probing to identify key iron-reducing microorganisms involved in anaerobic benzene degradation. ISME J 1: 643–653. Leuthner B, Leutwein C, et al. (1998) Biochemical and genetic characterization of benzylsuccinate synthase from Thauera aromatica: a new glycyl radical enzyme catalysing the first step in anaerobic toluene metabolism. Mol Microbiol 28(3): 615–628. Lovley D, Lonergan DJ (1990) Anaerobic oxidation of toluene, phenol, and para-cresol by the dissimilatory iron-reducing organism, GS-15. Appl Environ Microbiol 56(6): 1858–1864. Lovley DR, Woodward JC, et al. (1994) Stimulated anoxic biodegradation of aromatic-hydrocarbons using Fe(III) ligands. Nature 370(6485): 128–131.

Hydrocarbon Degradation Coupled to Metal Reduction Lovley DR, Woodward JC, et al. (1996) Rapid anaerobic benzene degradation with a variety of chelated Fe(III) forms. Appl Environ Microbiol 62(1): 288–291. McMaster J, Enemark JH (1998) The active sites of molybdenum- and tungsten-containing enzymes. Curr Opin Chem Biol 2(2): 201–207. Mobitz H, Friedrich T, et al. (2004) Substrate binding and reduction of benzoyl-CoA reductase: evidence for nucleotide-dependent conformational changes. Biochemistry 43(5): 1376–1385. Peters F, Rother M, et al. (2004) Selenocysteinecontaining proteins in anaerobic benzoate metabolism of Desulfococcus multivorans. J Bacteriol 186(7): 2156–2163. Peters F, Shinoda Y, et al. (2007) Cyclohexa-1,5-diene-1carbonyl-coenzyme A (CoA) hydratases of Geobacter metallireducens and Syntrophus aciditrophicus: Evidence for a common benzoyl-CoA degradation

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pathway in facultative and strict anaerobes. J Bacteriol 189(3): 1055–1060. Rooney-Varga JN, Anderson RT, et al. (1999) Microbial communities associated with anaerobic benzene mineralization in a petroleum-contaminated aquifer. Appl Environ Microbiol 65: 3056–3063. Schuhle K, Gescher J, et al. (2003) Benzoate-coenzyme A ligase from Thauera aromatica: an enzyme acting in anaerobic and aerobic pathways. J Bacteriol 185(16): 4920–4929. Stojanowic A, Mander GJ, et al. (2003) Physiological role of the F420-non-reducing hydrogenase (Mvh) from Methanothermobacter marburgensis. Arch Microbiol 180(3): 194–203. Wischgoll S, Heintz D, et al. (2005) Gene clusters involved in anaerobic benzoate degradation of Geobacter metallireducens. Mol Microbiol 58(5): 1238–1252.

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12 Anaerobic Degradation of Isoprene-Derived Compounds J. Harder Department of Microbiology, Max Planck Institute for Marine Microbiology, Bremen, Germany [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 958

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Molecular Oxygen and Anaerobic Catabolisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 958

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Isoprene and Branched Model Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 959

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Monoterpenes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 959

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Phytol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 960

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Squalene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 960

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Cholesterol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 960

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Bile Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 960

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Isoprenoids and Organic Matter Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 960

10 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 961

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_68, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Only a few of over 30,000 isoprenoic compounds have served as single carbon source in the isolation of anaerobic bacteria. Monoterpenes, phytol, cholesterol and oestradiol yielded denitrifying bacteria belonging to Castellaniella, Thauera, Sterolibacterium and Denitratisoma. The characterization of the special biochemistry of isoprenoid mineralization in these organisms has started.

1

Introduction

Anaerobic microbiology as a research field started with investigations on rumen microorganisms and fermentations. The mineralization of hydrocarbons became of interest since the 1980s (Grbic-Galic and Vogel, 1987). With a major focus on compounds of environmental concern, only few metabolite and isolation studies have addressed the transformation and mineralization of isoprenoid compounds. A second focus of anaerobic isoprenoid microbiology is the biotransformation of steroids in gut systems (Hylemon and Harder, 1999).

2

Molecular Oxygen and Anaerobic Catabolisms

Hydrocarbons including alkanes, alkenes and aromatics are slowly degraded due to physical and chemical characteristics. Oxygen-respiring microorganisms in culture have usually mono- or dioxygenases that utilize molecular oxygen as cosubstrate for the initial activation reactions of the hydrocarbons to alcohols. The world of anaerobic organisms starts when the oxygen concentration is insufficient to allow thermodynamically the mineralization of organic matter coupled to energy conservation. This threshold concentration varies, the cytochrome C oxidase has a higher oxygen threshold than the alternative oxidases that utilize ubiquinone as electron donor. Below the threshold and in dynamic systems with an oxygen flux, molecular oxygen is still present and available for reaction without energy conservation, e.g., Saccharomyces cerevisiae has a facultative fermentative metabolism, but proliferates only with a ribonucleotide reductase that is activated by the reaction of molecular oxygen with a dinuclear iron(II)center. The genome sequence has revealed the absence of an anaerobic ribonucleotide reductase (Harder and Follmann, 1990; Nordlund and Reichard, 2006). The anaerobic world can be divided in facultative anaerobes, mainly nitrate-, chlorate-, manganese(IV)- and iron(III)-respiring organisms living at a slightly positive redoxpotential and eventually with traces of molecular oxygen in dynamic open systems, and obligate anaerobes, mainly fermenting, sulfate-reducing and methanogenic organisms living in the absence of molecular oxygen at a negative redoxpotential of around 200 mV. Activation reactions of hydrocarbons in aerobic organisms without molecular oxygen are rarely observed, probably because the organisms are not competitive in the isolation procedures. The rediscovery of an oxygen-independent pathway in a denitrifying bacterium indicates the advantage of anaerobic enrichments for the identification of oxygen-independent pathways (Harder, 1997). Recently, the proteins of the anaerobic degradation pathway for cholesterol were found to be expressed also under oxic conditions (Chiang et al., 2008). This overview includes some of these molecular oxygen-independent pathways.

Anaerobic Degradation of Isoprene-Derived Compounds

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Isoprene and Branched Model Compounds

The biotransformation of isoprene under anaerobic conditions has not been reported. Isoprenoids have typically tertiary and quaternary carbon atoms. A tertiary carbon atom in a-position to a carboxylic CoA thioester does not hamper the b-oxidation pathway, typically a propionyl CoA is released. A tertiary carbon atom in b-position to a carboxylic CoA thioester requires additional reactions. The removal of the methyl group is best explored in the leucine biodegradation, the classical textbook example for the degradation of tertiary carbons. The b-hydroxyl-b-methylglutaryl CoA lyase transforms the tertiary carbon into a secondary and enables further b-oxidation. The feasibility of the mineralization of quaternary carbon atoms in anaerobic bacteria has been demonstrated by a number of denitrifying isolates, but biochemical studies have not been initiated (Kniemeyer et al., 1999; Probian et al., 2003).

4

Monoterpenes

Volatile organic compounds of plants are a significant part of the global carbon cycle. The monoterpenes present a major part and are well known as constituents of essential oils, e.g., pinene. Below toxic concentrations, the compounds may be mineralized by denitrifying microorganisms (Harder and Probian, 1995; Harder et al., 2000). The monoterpenedegrading Betaproteobacterium Alcaligenes defragrans has been renamed to Castellaniella defragrans (Foß et al., 1998; Ka¨mpfer et al., 2006). Initial metabolite studies have revealed the microbial capacity to transform monoterpenes and geranic acid has been identified as first ionic metabolite (Heyen and Harder, 1998, 2000). The degradation of linear monoterpenederived acids seems to be highly similar to the leucine degradation pathway. The biochemistry of the operons for acyclic terpene utilization (atu) and leucine/isovalerate utilization (liu) is currently intensively explored for Pseudomonas citronellolis and P. aeruginosa (Aguilar et al., 2008; Fo¨rster-Fromme et al., 2008).

Oxygenated monoterpenes are in general more toxic than non-functionalized monoterpenes due to the reactivity especially of the ketone and aldehyde groups (Harder et al., 2000). However, Thauera terpenica and Thauera linaloolentis could be isolated and initial studies revealed with an allylalcohol isomerase a novel enzyme activity (Foß and Harder 1997, 1998). Monoterpenes are not only mineralized by methanogenic cultures, but are also oxidized to cymene, p-isopropyl-methyl-benzene. The aromatization reaction is a typical diagenetic reaction of cyclohexene and cyclohexadiene compounds (Harder and Foß, 1999). Interestingly, cymene degrading denitrifying isolates are also growing on monoterpenes, e.g., limonene (Harms et al., 1999).

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Phytol

Phytol is a linear diterpene alcohol ubiquitous in nature as a side chain of chlorophyll. It is an important biogeochemical marker in organic geochemistry. A number of microbial transformation products have been identified and suggest a similar degradation pathway as discussed for acyclic monoterpene acids (Rontani and Volkman, 2003).

6

Squalene

The tail-to-tail condensation of two farnesyl diphosphates yields squalene, the precursor for all triterpenes. A marine gammaproteobacterium related to Marinobacter transformed the alkene by hydration to a diol with tertiary alcohols. A methylketone metabolite together with a carbon dioxide stimulation of the further degradation suggest the presence of a novel carbon– carbon cleavage reaction (Rontani et al., 2002).

7

Cholesterol

Denitrifying isolates, strain 72Chol (Harder and Probian, 1997) and the related Sterolibacterium denitrificans (Tarlera and Denner, 2003), completely mineralize cholesterol. A molecularoxygen independent hydroxylation reaction of the terminal tertiary carbon atom functionalizes the molecule, likely with a molybdenum-containing enzyme (Chiang et al., 2007). Twodimensional gel electrophoresis of 14C-labeled soluble proteins indicated the presence of the pathway also under oxic circumstances (Chiang et al., 2008). For other steroids, including hormones, the isolation of Denitratisoma oestradiolicum on 17b-oestradiol demonstrated the anaerobic biodegradation of the female sex hormone (Fahrbach et al., 2006). Enrichment cultures were also established for methanogenic, sulfateand iron-reducing conditions (Czajka and Londry, 2006).

8

Bile Acids

Cholate and deoxycholate circle in the enterohepatic cycle supporting the uptake of waterinsoluble nutrients. During the passage through the intestinal tract, cholate is dehydroxylated in the 7a-position by fermenting bacteria (Hylemon and Harder, 1999; Ridlon et al., 2006). A partial cholate oxidation under denitrifying conditions yielded 7,12-dihydroxy-1,4androstandiene-3,17-dione, but a complete mineralization has not been described (Philipp et al., 2006).

9

Isoprenoids and Organic Matter Degradation

The molecular biochemistry focuses usually on one substrate and one organism. In the environment, the degradation processes in sediments involves many compounds as substrates at the same time. Microcosm experiments coupled to an in-depth analysis of organic molecules can reveal the action of microbial communities on isoprenoid compounds in relation to

Anaerobic Degradation of Isoprene-Derived Compounds

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the chemical structure. Grossi et al. (2001) reported the anaerobic biodegradation of the microalga Nannochloropsis salina: phytol belonged to the most labile lipid constituents, whereas sterols exhibited the lowest loss. Within the sterols, cholesterol was more degraded than C29 sterols. Such mixed substrate studies are an opportunity to expand our knowledge on the microbial transformation and mineralization of isoprenoid compounds.

10

Research Needs

Of the over 30,000 described isoprene-derived natural substances, only a few have served as substrate in the enrichment and isolation of anaerobic microorganisms. The isolation of denitrifying bacteria on model substances has shown the existence of anaerobic mineralization pathways. In the future, the interest in new biochemical reactions will maintain the exploration of the biochemical pathways in these organisms.

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Fo¨rster-Frommne K, Chattopadhyay A, Jendrossek D (2008) Biochemical characterization of AtuD from Pseudomonas aeruginosa, the first member of a new subgroup of acyl-CoA dehydrogenases with specificity for citronellyl-CoA. Microbiology 154: 789–796. Grbic-Galic D, Vogel T (1987) Transformation of toluene and benzene by mixed methanogenic cultures. Appl Environ Microbiol 53: 254–260. Grossi V, Blokker P, Damste JSS (2001) Anaerobic biodegradation of lipids of the marine microalga Nannochloropsis salina. Org Geochem 32: 795–808. Harder J (1997) Anaerobic degradation of cyclohexane1,2-diol by a new Azoarcus species. Arch Microbiol 168: 199–204. Harder J, Follmann H (1990) Identification of a free radical and oxygen dependence of ribonucleotide reductase in yeast. Free Radic Res Commun 10: 281–286. Harder J, Foß S (1999) Anaerobic formation of the aromatic hydrocarbon p-cymene from monoterpenes by methanogenic enrichment cultures. Geomicrobiol J 16: 295–305. Harder J, Heyen U, Probian C, Foß S (2000) Anaerobic utilization of essential oils by denitrifying bacteria. Biodegradation 11: 55–63. Harder J, Probian P (1995) Microbial degradation of monoterpenes in the absence of molecular oxygen. Appl Environ Microbiol 61: 3804–3808. Harder J, Probian C (1997) Anaerobic mineralisation of cholesterol by a novel type of denitrifying bacterium. Arch Microbiol 167: 269–274. Harms G, Rabus R, Widdel F (1999) Anaerobic oxidation of the aromatic plant hydrocarbon p-cymene by

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newly isolated denitrifying bacteria. Arch Microbiol 172: 303–312. Heyen U, Harder J (1998) Cometabolic isoterpinolene formation from isolimonene by denitrifying Alcaligenes defragrans. FEMS Microbiol Lett 169: 67–71. Heyen U, Harder J (2000) Geranic acid formation, an initial reaction of anaerobic monoterpene metabolism in denitrifying Alcaligenes defragrans. Appl Environ Microbiol 66: 3004–3009. Hylemon PB, Harder J (1999) Biotransformation of monoterpenes, bile acids, and other isoprenoids in anaerobic ecosystems. FEMS Microbiol Rev 22: 475–488. Ka¨mpfer P, Denger K, Cook AM, Lee ST, Jackel U, Denner EBM, Busse HJ (2006) Castellaniella gen. nov., to accommodate the phylogenetic lineage of Alcaligenes defragrans, and proposal of Castellaniella defragrans gen. nov., comb. nov and Castellaniella denitrificans sp nov. Int J Syst Evol Microbiol 56: 815–819. Kniemeyer O, Probian C, Rossello´-Mora R, Harder J (1999) Anaerobic mineralization of quaternary carbon atoms: isolation of denitrifying bacteria on dimethylmalonate. Appl Environ Microbiol 65: 3319–3324.

Nordlund P, Reichard P (2006) Ribonucleotide reductases. Ann Rev Biochem 75: 681–706. Philipp B, Erdbrink H, Suter MJF, Schink B (2006) Degradation of and sensitivity to cholate in Pseudomonas sp. strain Chol1. Arch Microbiol 185: 192–201. Probian C, Wu¨lfing A, Harder J (2003) Anaerobic mineralization of quaternary carbon atoms: isolation of denitrifying strains on pivalic acid (2,2dimethylpropionic acid). Appl Environ Microbiol 69: 1866–1870. Ridlon JM, Kang DJ, Hylemon PB (2006). Bile salt biotransformations by human intestinal bacteria. J Lipid Res 47: 241–259. Rontani JF, Mouzdahir A, Michotey V, Bonin P (2002) Aerobic and anaerobic metabolism of squalene by a denitrifying bacterium isolated from marine sediment. Arch Microbiol 178: 279–287. Rontani JF, Volkman JK (2003) Phytol degradation products as biogeochemical tracers in aquatic environments. Org Geochem 34: 1–35. Tarlera S, Denner EBM (2003) Sterolibacterium denitrificans gen. nov., sp nov., a novel cholesteroloxidizing, denitrifying member of the bProteobacteria. Int J Syst Evol Microbiol 53: 1085–1091.

13 Degradation of Long-Chain Fatty Acids by SulfateReducing and Methanogenic Communities D. Z. Sousa1,2 . M. Balk1 . M. Alves2 . B. Schink3 . M. J. McInerney4 H. Smidt1 . C. M. Plugge1 . A. J. M. Stams1 1 Laboratory of Microbiology, Wageningen University, Wageningen, The Netherlands 2 Institute for Biotechnology and Bioengineering, Centre of Biological Engineering, University of Minho, Braga, Portugal 3 Department of Biology, University of Konstanz, Konstanz, Germany 4 Department of Botany and Microbiology, University of Oklahoma, Norman, OK, USA [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 964 2 General Biochemical Pathway of LCFA Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 964 3 Hydrogenation of Unsaturated Fatty Acids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 966 4 LCFA Degradation by Sulfate-Reducing Communities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 966 5 LCFA Degradation by Methanogenic Communities . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 967 6 Biogas Formation from LCFA Containing Waste Materials . . . . . . . . . . . . . . . . . . . . . . . . 969 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 975

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_69, # Springer-Verlag Berlin Heidelberg, 2010

964

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Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

Abstract: Lipids and long chain fatty acids (LCFA) are energy-rich compounds that support the growth of microorganisms that thrive in environments with a low redox potential. This chapter describes the properties of LCFA-degrading sulfate-reducing bacteria and LCFA-degrading methanogenic communities. The methanogenic conversion of LCFA requires syntrophic communities of acetogenic bacteria and methanogenic archaea. In this syntrophic cooperation, interspecies hydrogen transfer plays a key role. The understanding of the microbial interactions involved in LCFA degradation is essential to optimize the methane formation from LCFAcontaining waste streams in bioreactors. Sulfate-reducing and methanogenic communities degrade LCFA by b-oxidation, but the differences and similarities between the degradation of saturated and unsaturated LCFA are not yet fully understood. Generally, bacteria that degrade unsaturated fatty acids degrade saturated fatty acids also, but the opposite does not always seem to be the case.

1

Introduction

Lipids, natural oils, and fats are abundantly present in nature. Lipids are constituents of membranes of bacteria, archaea, and eukaryotes, while oils and fats are storage compounds for carbon and energy in all kinds of living organisms. Lipids are mostly esters of glycerol and long-chain fatty acids (LCFA) or, in the case of archaea, ethers of glycerol and long-chain alcohols. A substantial part of the dry weight of biomass is lipids, oils, and fats. These energyrich compounds can be anaerobically degraded by methanogenic and sulfate-reducing communities. Methanogens and sulfate-reducing bacteria are not known to hydrolyze lipids. This suggests that other anaerobic microorganisms are responsible for the synthesis and excretion of lipases for the initial attack of lipids. A variety of sulfate-reducing bacteria is able to degrade LCFA coupled to the reduction of sulfate. However, methanogens do not degrade LCFA. In methanogenic environments, LCFA are degraded by proton-reducing acetogenic bacteria to acetate and hydrogen, which are substrates for the methanogens. For energetic reasons, the growth of proton-reducing acetogenic bacteria on LCFA is possible only if acetate and, in particular, hydrogen are efficiently removed by methanogens. This results in an obligately syntrophic growth driven by interspecies hydrogen transfer (Schink, 1997; Schink and Stams, 2006). In this chapter, we present information on the physiology of sulfate-reducing bacteria and syntrophic methanogenic communities that are able to grow on LCFA.

2

General Biochemical Pathway of LCFA Degradation

LCFA degradation pathways have not been studied extensively in methanogenic and sulfatereducing communities at a biochemical or genetic level. Experiments with 14C-labeled LCFA indicated that degradation occurs by b-oxidation (Nuck and Federle, 1996; Weng and Jeris, 1976). Detailed studies on LCFA metabolism have been done in model organisms like Escherichia coli (DiRusso et al., 1999). LCFA biodegradation occurs through sequential steps: (1) LCFA adsorption to the cell surface, (2) LCFA uptake, and (3) LCFA conversion to lower molecular weight components via b-oxidation. A scheme of the b-oxidation cycle is shown in > Fig. 1. The end product in this cycle is acetyl-CoA.

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

13

. Figure 1 The b-oxidation pathway involved in LCFA degradation in sulfate-reducing bacteria, and acetogenic bacteria that grow in syntrophy with methanogens.

E. coli possesses a metabolic pathway to degrade LCFA anaerobically in the presence of nitrate, fumarate, or trimethylamine-N-oxide that is distinct from the Fad enzymes used for aerobic LCFA metabolism (Campbell et al., 2003; Klein et al., 1971). FadIJ, like FadAB (Pramanik et al., 1979), appear to form a multienzyme complex that contains enoyl-CoA hydratase, 3-hydroxyacyl-CoA dehydrogenase, and 3-oxoacyl-CoA thiolase activities (Campbell et al., 2003; Snell et al., 2002). In contrast to the gene organization in E. coli, genomic analyses of Syntrophus aciditrophicus (McInerney et al., 2007) and Syntrophomonas wolfei (http://www. jgi.doe.gov) indicate that separate genes encode for enoyl-CoA hydratase, 3-hydroxyacylCoA dehydrogenase, and 3-oxoacyl-CoA thiolase activities. Both the S. aciditrophicus and S. wolfei genomes contain multiple homologs encoding not only for enoyl-CoA hydratase, 3-hydroxyacyl-CoA dehydrogenase, and 3-oxoacyl-CoA thiolase activities but also for acetylCoA synthetase (AMP-forming) and acyl-CoA dehydrogenase activities (> Table 1). Presumably, the homologs differ in chain-length or substrate (saturated vs. unsaturated fatty acids) specificity. In sulfate-reducing bacteria that degrade LCFA completely to CO2, acetyl-CoA is further degraded via the acetyl-CoA cleavage pathway or a modified citric acid cycle (Schauder et al., 1986). Acetyl-CoA can also be converted to acetate as is the case for LCFA-degrading bacteria in methanogenic environments and in several sulfate-reducing bacteria. Acetate is then a substrate for acetoclastic methanogens (Methanosarcina and Methanosaeta) or acetate-utilizing sulfatereducing bacteria (Desulfobacter, Desulfobacterium, Desulforhabdus and Desulfobacca).

965

966

13

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

. Table 1 Homologues of b-oxidation genes in the genomes of syntrophic acetogenic bacteria Number of homologues Function

S. aciditrophicus

S. wolfei

Acyl-CoA synthetase (AMP-forming)

4

2

Acyl-CoA dehydrogenase

6

9

Enoyl-CoA hydratase

5

5

3-hydroxyacyl-CoA dehydrogenase

1

7

3-oxoacyl-CoA thiolase

2

5

Source: S. aciditrophicus genome (McInerney et al., 2007) and S. wolfei genome (http://www.jgi.doe.gov)

3

Hydrogenation of Unsaturated Fatty Acids

Some fermentative microorganisms are capable of hydrogenating unsaturated fatty acids, without being able to grow by these conversions (Mackie et al., 1991). Several anaerobic ruminal and human gut bacteria with biohydrogenation capabilities, belonging to the genera Butyrivibrio, Pseudobutyrivibrio, ‘‘Fusocillus,’’ Borrelia, Roseburia, and Clostridium, have been isolated and characterized (Devillard et al., 2007; Fukuda et al., 2005; Hunter et al., 1976; Kemp et al., 1984; Kopecny et al., 2003; Maia et al., 2007; Moon et al., 2008; Sachan & Davis, 1969; van de Vossenberg and Joblin, 2003; Wallace et al., 2006). Early studies suggested that the degradation of unsaturated LCFA, such as linoleic (C18:2) and oleic (C18:1) acids, proceeded via b-oxidation only after chain saturation (Novak and Carlson, 1970; Weng and Jeris, 1976). Canovas-Dias et al. (1991) detected palmitoleate (C16:1) as a transient product during oleate (C18:1) degradation, suggesting the occurrence of one b-oxidation step without prior chain saturation. Thus far, experimental data are lacking to define the exact pathway involved in the degradation of unsaturated LCFA. Nevertheless, an analysis of the free energy variation of different possible reactions involved in LCFA degradation can provide some insight into which pathways are most likely as illustrated for oleate in (> Table 2). The hydrogenation step is thermodynamically favorable as indicated by the Gibbs free energy change of oleate (C18:1) conversion to stearate (C18:0). The direct b-oxidation of oleate (C18:1) to palmitoleate (C16:1) is thermodynamically unfavorable under standard conditions (DG00 > 0). Oleate (C18:1) degradation by a combined hydrogenation and b-oxidation to form palmitate (C16:0) seems most likely, as the endergonic oxidation reaction could be driven by the exergonic hydrogenation reaction. In studies performed with anaerobic sludges, palmitate was indeed a main intermediate product in oleate degradation (Lalman and Bagley, 2001; Pereira et al., 2002).

4

LCFA Degradation by Sulfate-Reducing Communities

Baars (1930) described Vibrio ru¨bentschickii, a sulfate-reducing bacterium that is able to grow on LCFA and short-chain fatty acids. However, it never became really clear whether or not this culture was a pure culture of a sulfate-reducing bacterium (Postgate and Campbell, 1966).

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

13

. Table 2 Gibbs free energy changes (at standard conditions) for hydrogenation and b-oxidation reactions (presumably) involved in conversion of unsaturated fatty acids Reaction

DG00 (kJ/reaction)

Hydrogenation

Oleate-(C18:1) + H2 ! Stearate-(C18:0)

66

Oxidation

Oleate-(C18:1) ! Palmitoleate-(C16:1) + acetate- + 2 H2

+51

Hydrogenation + oxidation

Oleate-(C18:1) ! Palmitate-(C16:0) + acetate- + H2 + H+

15

Note: Values calculated with data from Thauer et al. (1977) and Mavrovpuniotis (1991)

Later, the ability of pure cultures of sulfate-reducing bacteria to grow on LCFA and short-chain fatty acids was shown by Widdel (1980). Currently, representatives of several genera of sulfate-reducing microorganisms are known to be able to grow on LCFA (> Fig. 2). > Table 3 lists some properties of sulfate-reducing bacteria that can grow on LCFA. Some of the sulfate-reducing bacteria that have been tested grow also on acetate, indicating that these sulfate-reducing bacteria degrade LCFA completely to CO2. However, many LCFAdegrading sulfate-reducing bacteria produce acetate as a side product.

5

LCFA Degradation by Methanogenic Communities

Syntrophomonas sapovorans was the first-described LCFA-degrading bacterium that grows in syntrophy with methanogens (Roy et al., 1986). The first-defined butyrate-degrading culture, consisting of Syntrophomonas wolfei and Methanospirillum hungatei, was described earlier by McInerney et al. (1981). This bacterium is able to degrade fatty acids with a chain length of up to 8 carbon atoms. Syntrophomonas sapovorans grows on LCFA with more than 12 and up to 18 carbon atoms and is able to utilize unsaturated LCFA, such as oleate (C18:1) and linoleate (C18:2). To date, 14 syntrophic LCFA-degrading microorganisms have been obtained in pure culture or in coculture with hydrogen-consuming microorganisms (> Table 4; > Fig. 2). All these acetogenic bacteria are capable of anaerobically degrading fatty acids with more than 4 carbon atoms and up to 18 carbon atoms . They all belong to the families Syntrophomonadacea (McInerney, 1992; Zhao et al., 1993; Wu et al., 2006a) and Syntrophaceae (Jackson et al., 1999). LCFA higher than lauric acid (i.e., with more than 12 carbon atoms) are utilized only by Syntrophomonas sapovorans, S. saponavida, S. curvata, S. zehnderi, S. palmitatica, Thermosyntrophica lipolytica, and Syntrophus aciditrophicus. T. lipolytica also grows syntrophically with methanogens on lipids such as olive oil, utilizing only the liberated fatty acid moieties and releasing the glycerol. Two lipases are excreted by this bacterium (Salameh and Wiegel, 2007). The LCFA degradation in methanogenic environments requires the syntrophic cooperation of LCFA-degrading bacteria and methanogens. Interspecies hydrogen transfer is a key process in methanogenesis. In 1967, Bryant and coworkers described for the first time an obligate syntrophic association between two microbial species (Bryant et al., 1967). They discovered that Methanobacillus omelianskii, originally believed to be a pure culture, (Stadtman and Barker, 1949; Barker, 1956) was not axenic. It was actually a coculture of a bacterium fermenting ethanol to acetate and H2 (the ‘‘S-organism’’), and an archaeon (later named

967

968

13

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

. Figure 2 (Continued)

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

13

Methanobacterium bryantii; Balch et al., 1979) that uses H2 to reduce carbon dioxide to methane. The conversion of ethanol to acetate and H2 by the ‘‘S-organism’’ was thermodynamically possible only in the presence of M. bryantii, which keeps the H2 concentration low. The concept of interspecies hydrogen transfer originating from these observations was fundamental to understand how compounds such as propionate, butyrate, and LCFA are degraded in methanogenic environments. > Table 5 summarizes the reactions involved in syntrophic LCFA degradation. To sustain their metabolism, LCFA-degrading bacteria have to couple the oxidation of NADH and FADH2 to proton reduction, which is energetically difficult. The midpoint redox potential of the couple H+/H2 at pH 7 is 414 mV, whereas the midpoint redox potential of the couples of NAD+/NADH and FAD/FADH2 are 320 mV and 220 mV respectively (Thauer et al., 1977). The DG00 values of the reactions NADH + H+ ! NAD+ + H2 and FADH2 ! FAD + H2 are about 18 and 38 kJ respectively. Methanogens are able to create a hydrogen partial pressure as low as 1 Pa. Under these conditions, the DG0 values of NADH and FADH2 oxidation are approximately 11 and + 9 kJ respectively. Thus, NADH oxidation, but not FADH2 oxidation, is feasible by the creation of a low hydrogen partial pressure by methanogens. It is not clear how LCFA-degrading bacteria solve the energetic problem of FADH2 oxidation coupled to hydrogen formation. Likely, a reversed electron transport mechanism is involved. The sequence analysis of the genes near fadK on the E. coli chromosome provides a clue about the nature of the reverse electron transport system involved in the syntrophic metabolism of LCFA. fadK mutants are impaired in anaerobic growth with fatty acids, and FadK is a putative acyl-CoA sythetase (Campbell et al., 2003). Genes ydiQRST are clustered with fadK on the E. coli chromosome, and ydiQRST have high sequence homology to fixABCX involved in the anaerobic carnitine metabolism in E. coli (Buchet et al., 1998; Walt and Kahn, 2002) and the reverse electron transport in N2-fixing bacteria (Earl et al., 1987; Edgren and Nordlund, 2004, 2006; Lindblad et al., 1996). The S. wolfei genome contains homologs to fixABCX, while S. aciditrophicus apparently uses another system for reverse electron transport (McInerney et al., 2007). Further research is needed to unravel the biochemistry of reverse electron transport.

6

Biogas Formation from LCFA Containing Waste Materials

LCFA are energy-rich compounds that are abundantly present in raw and waste materials (> Table 6). Thus, biogas formation from LCFA-containing waste represents a sustainable technology. About 1 m3 methane can be produced from 1 kg of LCFA. Lipids and LCFA are present in domestic and industrial wastewaters. In domestic wastewater, generally, lipids represent 20–25% of the total organic matter with concentrations ranging from 40 to 100 mg/L (Que´meneur and Marty, 1994). Lipids/LCFA concentrations in industrial wastewaters are more variable and highly dependent of the industrial process. Concentrations of 11.2–22.4 g lipids/L were found in an industrial wastewater from a wool-scouring process (Becker et al., . Figure 2 Phylogenetic tree of bacterial 16S rRNA gene sequences of the fatty-acid degrading sulfatereducing and acetogenic bacteria described in the > Tables 3 and > 4 (fatty-acid degrading bacteria are shown in bold). The tree was based on 16S rRNA gene sequences and calculated using the ARB software package (Ludwig et al., 2004). Thermotoga lettingae (AF355615) was used as outgroup.

969

0.68

0.6

0.45

1–1.5

0.5–0.7

1–1.5

0.8

2.3–3.0

1.0–1.3

Desulfatibacillum aliphaticivorans

Desulfatiferula olefinivorans

Desulfobacterium autotrophicum

Desulfocella halophila

Desulfococcus multivorans

Desulfofrigus fragile

Desulfonema limicola

Desulforegula conservatrix

Cell width (mm)

Desulfatibacillum alkenivorans

Cell length (mm)

2.6–3.0

2.0–5.0

3.2–4.2

1.5–2.5

2.0–4.0

1.5–2.5

0.8–5.0

2.2–5.5

1.2–4.5

Motility

Gram reaction

Spore formation 41.4 6.6–7.8/7.5

37.6 6.5–7.5/ND

5.8–7.6/ 6.5–7.3

57.4 6.7–7.6/7.3

35

47.6 ND/6.7

ND 52.1 ND/7.0–7.4







ND 45.5 6.6–8.3/7.5





DNA G + C content (mol%)

 +

 ND

ND

 gliding ND 34.5 6.5–8.8/7.6

 

 

 +

 +

 +

 

 

Growth pH (range/optimum)

ND/25–30

15–36/30

1.8–27/18

?/35

14–37/34

ND/25–28

16–38/ 30–36

15–40/ 28–35

20–40/ 28–34

Crotonate

Butyrate, C4:0

(+)

ND 

ND +

ND 

ND (+)

ND 

ND +

+

+

Caproate, C6:0

+

+

+

+

+

+

+

+

+

+

+

+

(+) +

+

ND +

Acetate, C2:0

ND 

+

+

Caprylate, C8:0 +



+

+

+

+

Caprate, C10:0

+

+

+

+

ND +

+

+

+

+

+

+

Laurate, C12:0

Myristate, C14:0 +

+

+



Palmitate, C16:0

+

+

ND +

+

+

+



Stearate, C18:0 +

+

+

+

CravoLaureau et al. (2007b)

CravoLaureau et al. (2004a)

ND ND CravoLaureau et al. (2004b)

Oleate, C18:1

ND ND ND Widdel (1980)

ND ND ND Brandt et al. (1999)

ND ND ND Brysch et al. (1987)

+

+



Linoleate, C18:2

+

+

+

+

+



ND ND ND Rees and Patel (2001)

ND ND ND Widdel et al. (1983)

ND ND ND ND ND ND Knoblauch et al. (1999)

+

+

+

+

+



13

Growth temperature (range/optimum)( C)

. Table 3 Selected characteristics of fatty-acid degrading sulfate-reducing bacteria

References

970 Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

1.0–1.3

0.8–1.2

0.7–0.8

0.5–0.7

0.5–0.7

0.8–1.0

1

0.5–0.8

1.2–2.0

Desulfatirhabdium butyrativorans

Desulfosarcina cetonica

Desulfospira joergensenii

Desulfotignum balticum

Desulfoarculus baarsii

Desulfothermus naphthae strain TD3

Thermodesulforhabdus norvegicus

Desulfotomaculum geothermicum

Desulfotomaculum sapomandens

5–7

2.3

205

2.0–3.5

1.5–4.0

1.5–3.0

1.0–2.0

1.8–2.7

2.6–3.5

1.5–2.5

 +

 +

 +

 ND

 +

 +

 

 

 

 

+

+















 6.7–9.0/7.4

6.5–8.2/7.3

6.5–8.2/7.3

6.5–7.9/ 7.0–7.4

ND/7.0

6.1–7.7/6.9

48

6.3–8.5/7.0

50.4 6–8/7.2–7.4

51

37.4 6.1–7.1/ 6.5–6.8

66

62

50

59

55.1 6.5–8.0/7.0

51

+, weak growth; , motility not clear; ND, not determined or not reported

1.0–1.5

Desulfosarcina variabilis

20–43/38

37–56/54

44–74/60

50–69/ 60–65

20–39/ 35–39

10–42/ 28–32

8.0–30/ 26–30

4–37/30

15–37/ 28–30

15–38/33 (+)

(+)



ND +

ND 

ND +

ND 

ND +

+

+

ND +

+

ND (+)

+

+

+

+

+

+

+

+

+

+

+

+

+

+



+

+

+

+

+

+

+

+

+

+

+

ND +

+

ND ND +

+

 Widdel (1980)

ND ND Balk et al. (2008)



ND ND ND Galushko and Rozanova (1991)

+



+

+

+

+

+

+

+

ND ND CordRuwisch and Garcia (1985)

ND ND Daumas et al. 1988)

ND ND Beeder et al. (1995)

ND ND Rueter et al. (1994) and Jakobsen et al. (2006)

ND ND Kuever et al. (2005)

ND ND Kuever et al. (2001)

ND ND ND ND Finster et al. (1997)

+

ND +

ND +

ND +

+

+

+

+

ND ND ND ND +

+

+

+

+

ND +

+

ND ND +



+

ND ND ND ND ND +

+

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

13 971

0.4

0.4– 0.5

0.5– 0.7

0.6– 0.9

Syntrophomonas cellicola

Syntrophomonas curvata

Syntrophomonas erecta erecta

Cell width (mm)

Syntrophomonas bryantii

Cell length (mm)

2.0– 8.0

2.3– 4.0

3.0– 10.0

4.5– 6.0

Spore formation

Motility

Gram reaction

 +

 +

 +

DNA G + C content (mol%)

48.8 6.5–8.5/ 7.0–7.5

37.6 6.5–7.5/ ND

Growth pH (range/optimum)

 43.2 6.0–8.8/ 7.8

 46.6 6.3–8.4/ 7.5

+

  +

Growth temperature (range/optimum)( C) 25–47/ 37–40

20–42/ 35–37

25–45/ 37–40

20–40/ 28–34

Crotonate +

+

+

+

Butyrate + pentenoate +

+

Butyrate, C4:0

+



ND 

+

+

+

ND ND +

Butyrate + DMSO

ND ND 



Caproate, C6:0 +

+

+

+

Caprylate, C8:0 +

+

+

+

Caproute, C10:0 

+



+

Laurate, C12:0 

+





Myristate, C14:0 

+





Palmitate, C16:0 

Stearate, C18:0

ND ND Methanospirilum hungatei Desulfovibrio sp. E70



+



+



+





Methanospirillum hungatei

Methanobacterium formicicum

ND ND ND ND Methanobacterium formicicum



Oleate, C18:1

Substrate utilization in co-culture with a syntrophic partner

Linoleate, C18:2

Zhang et al. (2005) and Wu et al. (2006b)

Zhang et al. (2005) and Zhang et al. (2004)

Wu et al. (2006a)

Zhao et al. (1990), Stieb and Schink (1985) , and Wu et al. (2006a)

13

Syntrophic partner used

Substrate utilization in pure culture

Acetate, C2:0

. Table 4 Selected characteristics of fatty-acid oxidizing acetogenic bacteria that grow in syntrophy with methanogens

References

972 Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

0.5– 0.7

0.4– 0.6

0.4– 0.6

0.5

0.4– 0.5

0.5– 1.0

0.4– 0.7

0.5– 0.7

Syntrophomonas erecta sporosyntropha

Syntrophomonas palmitatica

Syntrophomonas saponavida

Syntrophomonas sapovorans

Syntrophomonas wolfei methylbutyratica

Syntrophomonas wolfei wolfei

Syntrophomonas zehnderi

Syntrophus aciditrophicus

1.0– 1.6

2.0– 4.0

2.0– 7.0

3.0– 6.0

2.5

2.0– 4.0

1.5– 4.0

4.0– 14.0

+

40.6 5.5–8.4/ 7.0

47.3 6.5–8.5/ 7.0–7.6

6.3–8.1/ 7.3

ND

+

ND

ND

 45.1 6.2–8.1/ 7.0–7.5

+

 ND

 ND

   43.1 ND

 +

 +



 +

 +

   45.0 6.5–8.0/ 7.0

 + +

+

+

+



25–42/35 +

+

ND +



ND 

ND ND 

ND 

+

+

+

+

ND ND ND +

25–40/37  ND ND 

25–45/ 35–37

25–45/ 37–40



+

ND ND ND +

+



25–45/35  

ND

30–50/37 +

20–48/ 37–40

+

+

+

+

+

+

+

+

+

+

+

+

+

+

+

+

+





+

+

+



+





+

+

+



+





+

+

+



ND ND ND +

+





+

+

+



+

+





+

+

+



+







Methanospirillum hungatei

Methanospirillum hungatei Desulfovibrio sp. G11

Methanobacterium formicicum

Methanobacterium formicicum





Methanobacterium formicicum

Methanospirillum hungatei Desulfovibrio sp. G11

ND ND Methanospirillum hungatei Desulfovibrio sp. G11

+



ND ND Methanobacterium formicicum

+







Jackson et al. (1999)

Sousa et al. (2007c)

McInerney et al. (1979), McInerney et al. (1981), and Zhang et al. (2005)

Wu et al. (2007)

Zhang et al. (2005) and Roy et al. (1986)

Lorowitz et al. (1989) and Wu et al. (2007)

Hatamoto et al. (2007)

Wu et al. (2006b)

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

13 973

0.3– 0.4

Thermosyntropha lipolytica

Cell length (mm)

2.0– 3.5

2.0– 4.0

Motility

Gram reaction

DNA G + C content (mol%)

Spore formation

 51.0 5.8–7.5/ 6.5–7.0

Growth pH (range/optimum)

–a   43.6 7.1–9.5/ 8.1–8.9

 +

Growth temperature (range/optimum)( C) 52–70/ 60–66

45– 60/55

Crotonate +

+

Acetate, C2:0

Butyrate + DMSO

Butyrate + pentenoate ND ND 

ND ND 

Butyrate, C4:0 +

+

Caproate, C6:0 +

+

Caprylate, C8:0 +

+

Caproute, C10:0 +

þ

Laurate, C12:0 +



Myristate, C14:0 +



+



Palmitate, C16:0

a Cells stain Gram-negative in both exponential and stationary phase, but the organism has a Gram-positive cell wall ultrastructure Substrate utilization, +, utilized, , poorly utilized, , not utilized, ND, not determined or not reported

0.4– 0.5

Cell width (mm)

Syntrophothermus lipocalidus

+



Stearate, C18:0

Substrate utilization in co-culture with a syntrophic partner

+



Oleate, C18:1

Substrate utilization in pure culture

+



Linoleate, C18:2

Methanobacterium strain JW/VSM29

Methanobacterium thermoautotrophicum

Svetlitshnyi et al. (1996)

Sekiguchi et al. (2000)

13

Syntrophic partner used

. Table 4 (Continued)

References

974 Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

13

. Table 5 Some reactions involved in LCFA degradation in methanogenic environments and the Gibbs free energy changes DG00 (kJ/reaction)

DG0 (kJ/reaction)

oleate- + 16H2O ! 9 acetate- + 15H2 + 8H+

+338

177

stearate- + 16H2O ! 9 acetate- + 16H2 + 8H+

+404

139

+353

124

+48

22

Fatty acids oxidation reactions

palmitate- + 14H2O ! 8 acetate- + 14H2 + 7H butyrate- + 2H2O ! 2 acetate- + 2H2 + H+

+

Methanogenic reactions 4H2 + HCO3 + H+ ! CH4 + 3 H2O

136

14

acetate- + H2O ! HCO3+ CH4

31

31

DG00 is the Gibbs free energy change under standard conditions (temperature of 25  C, solute concentrations of 1 M and gas partial pressure of 105 Pa); DG0 was calculated for fatty acids concentrations of 1 mM, considering acetate stoichiometric accumulation (9, 8 or 2 mM for oleate/stearate, palmitate and butyrate degradation, respectively); PH2 = 1 Pa; PCH4 = 104 Pa. Values of standard Gibbs energy of formation for the various compounds were obtained with data from Thauer et al. (1977) and Mavrovouniotis (1991).

1999). Also, a relatively high concentration of lipids, that is, 6.6 g/L, was measured in olive oil mill effluents (Beccari et al., 1998). Lower concentrations (0.4–1.7 g lipids/L) were detected in a sunflower oil mill wastewater with LCFA concentrations in the range 0.2–1.3 g/L (Saatci et al., 2003). Total lipids in dairy wastewaters range from 0.9 to 2.0 g/L (Kim et al., 2004). Wastewaters and waste streams that contain high concentrations of lipids and LCFA may yield high levels of methane in an anaerobic digestion process. A problem associated with anaerobic treatment of lipids and LCFA is their poor solubility. LCFA were found to be inhibitory for methanogens (Lalman and Bagley, 2001, 2002; Kim et al., 2004; Pereira et al., 2003, 2004). The inhibitory effects are reversible and are often associated with the interactions of lipids and LCFA with the cell wall, preventing the conversion of other compounds due to their physical interaction (Pereira et al., 2005). However, by applying a sequence of loading and digestion phases in anaerobic reactors, high rates of methanogenesis and complete methanogenesis could be obtained (Pereira et al., 2002, 2003, 2004). Recently, it was demonstrated that continuous high rate anaerobic treatment of LCFA was possible by applying a relatively short start-up period with three feeding and three feed less phases. Methane recovery of up to 72% was obtained when a bioreactor was fed with an organic loading rate of 21 kgCOD m3 day1 and retention time of 9h (Cavaleiro et al., 2009). These findings offer excellent prospects for a sustainable energy production from lipids and LCFA.

7

Research Needs

Numerous strains of LCFA-degrading sulfate-reducing bacteria and LCFA-degrading acetogenic bacteria that grow in syntrophy with methanogens have been isolated and characterized. Although these bacteria have been relatively well studied from the physiological point of view, biochemical and genetic studies of the pathways involved in LCFA degradation are scarce. The

975

Dairy wastewater

2.2

2.6

1.0

Beef tallow

Domestic sewage

1.4

27.0

16.4

28.1

21.0

21.0

Whole milk

Chicken fat

27.7

Cocoa butter

6.0

11.0 35.7

1.0

1.4

Cotton seed oil

42.9

Soybean oil

1.4

Palmitate, C16:0

Saturated-LCFA

Myristate, C14:0

14.3

7.0

Laurate, C12:0

Olive oil

Palm oil

Raw materials/ Wastewaters

7.0

8.1

20.0

4.3

6.0

32.9

2.9

4.8

2.4

4.8

Stearate, C18:0

0.9

3.8

6.7

2.0

0.5

1.0

1.4

0.7

Palmitolate, C16:1

37.0

30.5

37.6

42.4

39.0

33.8

15.2

21.9

71.4

39.0

13.0

29.2

2.9

20.0

13.0

4.3

51.9

49.0

5.5

10.0

Linoleate, C18:2

Unsaturated-LCFA Oleate, C18:1

Kim et al. (2004)

Que´merneur and Marty (1994)

Taylor (1965)

Taylor (1965)

Hanaki et al. (1981)

Taylor (1965)

Taylor (1965)

Taylor (1965)

Taylor (1965)

Taylor (1965)

References

13

. Table 6 Saturated- and unsaturated-LCFA present in raw materials and wastewaters (shown as % of total LCFA) (adapted from Hwu, 1997)

976 Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

Degradation of Long-Chain Fatty Acids by Sulfate-Reducing and Methanogenic Communities

13

genomes of Desulfatibacillum alkenivorans and the acetogenic bacterium Syntrophomonas zehnderi are presently being sequenced (US Department of Energy, Joint Genome Institute). Comparison of the genomes of these bacteria and the genomes of related bacteria that are not well able to grow on LCFA may provide insight into LCFA-degrading pathways and the regulation of these pathways. In addition, it may allow an elucidation of the differences and similarities between degradation of saturated and unsaturated LCFA. Studies with methanogenic communities have shown that bacteria that were enriched with unsaturated fatty acids are able to degrade a wide range of saturated fatty acids, but the opposite is not the case (Sousa et al., 2007a, b). As lipids and LCFA are highly energetic compounds, further research is needed to get insight into how such compounds can be efficiently and completely converted to biogas. The challenge is to implement the syntrophic nature of the microorganisms involved in the most appropriate reactor configuration and the most optimal process operation.

Acknowledgments Our research on anaerobic LCFA degradation was made possible by the grants of the Fundac¸a˜o para a Cieˆncia e Tecnologia (FCT) and the Fundo Social Europeu (FSE) (SFRH/BD/8726/ 2002), the Wageningen Institute for Environmental and Climate Research (WIMEK), the Darwin Center for Biogeology of Netherlands Organization for Scientific Research (NWO), the NWO divisions Earth and Life Sciences (ALW) and Chemical Sciences (CW), and the Technology Foundation STW.

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13

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Thauer RK, Jungermann K, Decker K (1977) Energy conservation in chemotropic anaerobic bacteria. Bacteriol Rev 41: 100–180. van de Vossenberg JLCM, Joblin KN (2003) Biohydrogenation of C18 unsaturated fatty acids to stearic acid by a strain of Butyrivibrio hungatei from the bovine rumen. Lett Appl Microbiol 37: 424–428. Wallace JR, Chaudhary LC, McKain N, McEwan NR, Richardson AJ, Vercoe PE, Walker ND, Paillard D (2006) Clostridium proteoclasticum: A ruminal bacterium that forms stearic acid from linoleic acid. FEMS Microbiol Lett 265: 195–201. Walt A, Kahn MJ (2002) The fixA and fixB genes are necessary for anaerobic carnitine reduction in Escherichia coli. J Bacteriol 184: 4044–4047. Weng C, Jeris JS (1976) Biochemical mechanisms in methane fermentation of glutamic and oleic acids. Water Res 10: 9–18. Widdel F (1980) Anaerober Abbau von Fettsa¨uren und Benzoesa¨ure durch neu Isolierte Arten Sulfatreduzierender Bakterien. Ph.D. thesis, Go¨ttingen University, Go¨ttingen. Widdel F, Kohring GW, Mayer F (1983) Studies on dissimilatory sulfate-reducing 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. Wu C, Liu X, Dong X (2006a) Syntrophomonas cellicola sp. nov., a novel spore-forming syntrophic bacterium isolated from a distilled-spirit-fermenting cellar and assignment of Syntrophospora bryantii to Syntrophomonas bryantii sp. nov., comb. nov. Int J Syst Evol Microbiol 56: 2331–2335. Wu C, Liu X, Dong X (2006b) Syntrophomonas erecta subsp. sporosyntropha subsp. nov., a spore-forming bacterium that degrades short chain fatty acids in co-culture with methanogens. Syst Appl Microbiol 29: 457–462. Wu CG, Dong XZ, Liu XL (2007) Syntrophomonas wolfei subsp. methylbutyratica subsp. nov., and assignment of Syntrophomonas wolfei subsp. saponavida to Syntrophomonas saponavida sp. nov comb. nov. Syst Appl Microbiol 30: 376–380. Zhang CY, Liu XL, Dong XZ (2004) Syntrophomonas curvata sp. nov., an anaerobe that degrades fatty acids in co-culture with methanogens. Int J Syst Evol Microbiol 54: 969–973. Zhang CY, Liu XL, Dong XZ (2005) Syntrophomonas erecta sp. nov., a novel anaerobe that syntrophically degrades short-chain fatty acids. Int J Syst Evol Microbiol 55: 799–803. Zhao H, Yang D, Woese CR, Bryant MP (1993) Assignment of fatty acid-b-oxidizing syntrophic bacteria to Syntrophomonadaceae fam. nov. on the basis of 16S rRNA sequence analysis. Int J Syst Bacteriol 43: 278–286.

Part 4

Enzymology

14 Diversity and Common Principles in Enzymatic Activation of Hydrocarbons F. Widdel* . F. Musat Max Planck Institute for Marine Microbiology, Bremen, Germany *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 984

2

Functionalization of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 985

3 3.1 3.2 3.3 3.4 3.5

Aerobic Activation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 986 Dioxygen as a Reactant . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 986 Some Common Principles of Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 988 Heme-Iron Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 990 Non-Heme Di-Iron Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 993 Particulate Methane Monooxygenase (Particulate Di-Metal or Tri-Metal Monooxygenase) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 995 3.6 Non-Heme Mono-Iron Oxygenases (Ring Hydoxylating Dioxygenases) . . . . . . . . . . . . 996 3.7 Flavin-Containing Monooxygenases, and Monooxygenases with Unknown Mechanism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 996 4 Anaerobic Activation of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 998 4.1 Anaerobic Activation of Methane . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 999 4.2 Glycyl Radical Enzymes for Anaerobic Activation of Non-Methane Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1001 4.3 Ethylbenzene Dehydrogenase, a Molybdo-Enzyme . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1002 4.4 Unknown and Hypothesized Reactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1003 5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1004

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_70, # Springer-Verlag Berlin Heidelberg, 2010

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Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

Abstract: Hydrocarbons are apolar compounds devoid of functional groups and therefore exhibit (with some exceptions) low chemical reactivity at room temperature. Utilization of hydrocarbons by microorganisms as growth substrates is initiated by the introduction of a functional group. An astounding diversity of activation reactions has evolved in microorganisms, notably in bacteria. Saturated hydrocarbons are activated by initial C–H-bond cleavage, while unsaturated (including aromatic) hydrocarbons are activated by an addition of a co-reactant to form an initial s-bonded adduct. There is a principal difference between co-reactants and activation reactions in (1) aerobic and (2) anaerobic microorganisms. (1) Aerobic microorganisms always make use of molecular oxygen as a co-substrate so as to introduce one or two oxygen atoms by means of oxygenases. These enzymes usually contain metals (iron, copper). A common principle is the reduction of metal-bound O2 to the peroxide level; this converts into a metal-bound oxygen atom that performs the primary attack on the hydrocarbon. (2) Mechanisms in anaerobic activation of hydrocarbons are principally different. The anaerobic oxidation of methane is associated with a redox reaction of a nickel cofactor that is also involved in methanogenesis. The apparently most widely employed anaerobic activation mechanism of non-methane alkanes and alkyl-substituted aromatic hydrocarbons is a C–H-bond cleavage by a protein-hosted radical followed by addition of the radical product to fumarate; this results in a substituted succinate. A few alkyl-substituted aromatic hydrocarbons may be anaerobically hydroxylated (with the HO-group originating from H2O) at the side chain. In addition, there may be yet unknown mechanisms in anaerobic hydrocarbon activation.

1

Introduction

Utilization of hydrocarbons as carbon sources and electron donors (viz. as organic substrates) for growth is a domain of microorganisms, in particular of bacteria. Whereas higher organisms may only partially oxidize (oxygenate) some hydrocarbons, microorganisms can fully degrade numerous hydrocarbons of all major groups, the alkanes, alkenes, alkynes and aromatic hydrocarbons. Such degradation may occur aerobically with O2, or anaerobically with nitrate, ferric iron, sulfate or other electron acceptors. Any attempt of an overview of the metabolism of hydrocarbons in microorganisms is therefore not only confronted with the chemical diversity of hydrocarbons and their reactivities, but also with various microbial life styles. The metabolic degradation of hydrocarbons by microorganisms is conventionally treated in separate study areas (aliphatic vs. aromatic hydrocarbons; aerobic vs. anaerobic degradation pathways; physiology and overall metabolic pathways vs. enzyme mechanisms and structures), often with limited exchange and synopsis. But despite such separate treatment, the study areas are sometimes dealing with related questions. A very central of these questions concerns the ‘‘metabolic challenge’’ to channel an apolar, unreactive compound composed only of carbon and hydrogen into the metabolism. The hydrocarbon must be functionalized. In the present brief overview, the various modes of hydrocarbon activation are primarily divided into aerobic and anaerobic mechanisms, with subdivision according to principal types of enzymes. Emphasis is on ‘‘synoptic views’’ and generalizations (as far as possible) rather than on all ‘‘variations of themes’’, and on the activation of true hydrocarbons (compounds composed only of H and C). Subsequent reactions (e.g., alcohol oxidation, catechol ring cleavage) or oxygenation of hydrocarbon-like compounds with polar groups

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

14

are not included or mentioned only briefly when needed in the larger context. Also the utilization of alkyne hydrocarbons as non-natural compounds is not included (for enzymatic acetylene hydratation see Seiffert et al., 2007).

2

Functionalization of Hydrocarbons

Two principles may be distinguished among the initial steps in biochemical hydrocarbon activation by living organisms, activation at (a) a saturated (sp3) and (b) an unsaturated (mostly sp2) carbon atom: (a) If activation occurs at a saturated (sp3) carbon atom, the reaction has to begin with an attack on the C–H-bond; attack on a C–C-bond has never been observed. C–H-bond cleavage leads to a truncated carbon atom as a transition state to which a polar moiety is added. The activation reaction is thus a substitution (C–H ! C–X). Homolytic C–Hbond cleavage, viz. formation of a carbon radical (C–H ! C +  H), is most common. The energy for homolytic C–H-bond cleavage at sp3 carbon atoms of various hydrocarbons (in the gas phase ca. between 360 and 440 kJ mol1), which determines the activation energy, depends on the carbon compound (for compilation of energies of homolytic C–H-bond cleavage see > Chapter 1 in this volume; Widdel et al., 2007). Homolytic cleavage is facilitated if the carbon radical is stabilized (delocalized) by an adjacent p-electron system (resonance stabilization). In special cases, also heterolytic C–H-bond cleavage seems to be possible. A hydride (H) may be removed if the resulting carbenium (>C+) ion gains enough stability via adjacent carbon atoms and a p-electron system. In this case, an HO-ion may be added subsequently (see ethylbenzene dehydrogenase, 4.3). In a unique case, the anaerobic activation of methane, the CH4 molecule may directly react with a high-valent nickel complex to form a metal-organic compound ([Ni3+]–CH3; see Section 4.1). (b) Double bonded carbon atoms or those of aromatic rings (sp2 carbon atoms) are less likely to be biochemically activated by an initial C–H-bond cleavage because of the high bond energy (around 470 kJ mol1). A primary nucleophilic attack is essentially excluded in the case of hydrocarbons. However, unsaturated hydrocarbons can be attacked by strong electrophilic co-reactants that lead to primary s-adducts (the attacked carbon atom converts from sp2 to sp3). The mechanism may involve either an electron pair or a radical. The primary adduct then leads to a stable product via intramolecular rearrangements or by reaction with a further reactant. In the case of aromatic rings (AR), formation of the stable product can result in a net substitution at one carbon atom (AR–H ! AR–OH). However, this must not be confused with a substitution via C–H-bond cleavage as in the case of saturated carbon atoms. Whether or not the ‘‘classical’’ electrophilic substitution at an aromatic ring with an electrophilic cation (AR–H + X+ ! AR–X + H+), a frequent principle in aromatic chemistry, plays a role in biological activation of aromatic hydrocarbons is a matter of discussion. Unsaturated hydrocarbons are also prone to an addition of radical species, a reaction type discussed for dioxygenation of aromatic rings. As in the case of saturated hydrocarbons, the radical contained in the primary activation product would then require ‘‘neutralization’’ by a second radical that also results from the net reaction.

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Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

Aerobic Activation of Hydrocarbons

Utilization of hydrocarbons in aerobic microorganisms is always initiated by enzymatic reaction with dioxygen as a co-substrate to introduce hydroxyl functions. From a chemical point of view, dioxygen is an ideal co-substrate to achieve activation of carbon compounds including hydrocarbons.

3.1

Dioxygen as a Reactant

Dioxygen has fascinated biochemists since decades. O2 is a strong chemical and biochemical oxidant from a thermodynamic and electrochemical point of view (O2/2H2O: E  = +1.23 V, E 0 = +0.818 V) that can, in principle, enable highly exergonic and exothermic incomplete or complete oxidation reactions of carbon compounds. For instance, a hypothetical addition of O2 to ethane to yield ethanediol (‘‘CH3–CH3 + O2 ! CH2OH–CH2OH’’) would have a DG  of 290 and a DH  of 372 per mol. However, normal dioxygen does not react at room temperature with the majority of organic compounds on a time scale of years. This striking ‘‘kinetic inhibition’’ (if, for instance, compared to the aggressiveness of Cl2 or Br2) is crucial for the existence of life and organic compounds on our oxic Earth and for the use of water as electron donor for oxygenic photosynthesis. The co-existence of organic compounds and O2 is explained by the unique electronic state of the element in its ground state. The O2 molecule has two electrons more than N2 (in which occupation of the highest bonding molecular orbitals is complete). For the two additional electrons in O2, only anti-bonding (symbol: *) molecular orbitals are available. In ground state (‘‘normal’’) O2, the two electrons necessarily (according to Hund’s rule) occupy separately the two lowest anti-bonding, energetically equivalent (degenerate) orbitals leading to triplet oxygen (3Sg O2), the paramagnetic diradical (> Fig. 1). A concerted combination of these electrons with those of organic compounds in their common singlet state is extremely unlikely (‘‘spin-forbidden’’; only electrons with anti-parallel spin can combine to form a new bond); with other words, normal dioxygen does not insert into or add to most organic compounds. For reaction with organic compounds, the electronic state of dioxygen has to change, yielding energized (activated) oxygen. This can be achieved (a) physically or (b) chemically. (a) Physical activation in nature occurs by quantum energy transfer from compounds (pigments, dyes, etc.) excited by absorption of daylight1; O2 itself is not excited by daylight. The most common of the two known excited states, the diamagnetic singlet oxygen (1Dg O2) with antiparallel electron spins in one antibonding orbital (> Fig. 1), has considerable reactivity and destructive potential. It behaves as a substance rather different from triplet oxygen, even though both O2 molecules have the same oxidation state (0). For instance, singlet oxygen spontaneously adds to double bonds or reacts with C–H-groups next to double bonds (allylic carbon groups) to form organic hydroperoxides (R–O–O–H). (b) Chemical activation of dioxygen occurs by partial fill-up of orbitals with electrons from donor molecules (reduction by electron transfer), or by addition to transition metal ions which in their incompletely filled d-orbitals also have unpaired electrons (Bugg, 2003). 1

Chemical generation of singlet dioxygen is difficult to achieve. An example is the oxidation of hydrogen peroxide with hypochlorous acid (H2O2 + HClO ! H2O + 1Dg O2 + Cl + H+).

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

14

. Figure 1 Occupation of individual bonding and antibonding (*) orbitals in molecules of dioxygen species (modified from Halliwell and Gutteridge, 1984). A second singlet state of O2 with higher energy is not shown here. The two atomic 2p orbitals along the connecting axis that form the s–bond between the nuclei are usually designated as 2px orbitals. This leaves 2py and 2pz to form the p-bonds. However, conventions may differ, and the atomic orbitals along the connecting axis may be also termed 2pz, leaving 2px and 2py for the p-bonds.

Triplet oxygen can take up electrons stepwise from electron donors with appropriate redox potential, for instance from polyphenolic compounds (such as hydroquinols), reduced flavins, ascorbate, thiols, certain carbanions, organic radicals, or unsaturated organic compounds excited by light. The first reduction step leads to superoxide (O2 ; > Fig. 1), a reaction with rather negative redox potential (O2/O2 , E  = 0.33 V for gaseous standard state of O2; Elstner, 1990; Sawyer, 19812). Such negative redox potential may explain why O2 can bind reversibly to hemoglobin in its native FeII-state, without net conversion to superoxide; the redox potential of FeII in hemoglobin is not negative enough, a property crucial for the existence of animal life. Superoxide can further react in three ways. (1) As a reductant, it can be reoxidized to groundstate O2. (2) Superoxide can combine with an organic radical to form an organic peroxide or organic hydroperoxide (R + O2 + H+ ! R–O–OH). Direct combination of superoxide with the organic radical resulting from O2 reduction is, at a glance, ‘‘spin-forbidden’’. Nevertheless, there must be possibilities for spin inversion. A prominent example is the oxygenase by-activity of ribulose-1,5-bisphosphate carboxylase which cleaves ribulose-1,5-bisphosphate by O2 into 2-phosphoglycolate and 3-phosphoglycerate. The reaction probably involves oxygen reduction to superoxide by an organic substrate cation, spin inversion, and combination

Another redox potential often indicated, E˚ = −0.16 V, is based on standard activity (concentration) of aqueous O2 (dissolved in H2O) dissolved. Despite the relatively negative standard redox potential, the low O2 concentration that is in equilibrium with O2 can nevertheless be relevant with respect to reactivity. Also, the equilibrium concentration of O2 (that is formed by a one-electron step) does not decrease as dramatically (factor 10 per 0.0592 V, according to Nernst equation) with increasing redox potential as that of species formed by a two electron step (factor 100 per 0.0592 V). The reduction around pH = 7 does not involve a proton, because superoxide is deprotonated (O2 /HO2 , pKa = 4.6). 2

987

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Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

to an organic peroxide that disintegrates into the two carbon compounds (Lorimer, 1981). (3) Superoxide can be further reduced to inorganic peroxide (O22; > Fig. 1); the free form of the latter is usually the fully protonated one, H2O2 (H2O2/HO2, pKa = 11.8). Hydrogen peroxide formation has a high redox potential (O2 + 2H+/ H2O2; E 0 = +0.89 V). Hydrogen peroxide reacts with several organic compounds that possess activated carbon atoms, for instance to yield organic hydroperoxides that can desintegrate. Further electron uptake by inorganic or organic peroxides leads to separation of the O–O-bond. If peroxide is further reduced by another one-electron step, the hydroxyl radical (HO ), one of the most reactive oxygen species, is generated. Also the Fenton reaction, which involves Fe2+ as reductant, is usually formulated with the hydroxyl radical as product (Fe2+ + H2O2 + H+ ! Fe3+ + HO + H2O). However, the prevalent form may be an iron(IV)-oxo (ferrylIV) species in equilibrium with the hydroxyl radical ([FeIII–OH]2+ +  OH Ð [FeIV=O]2+ + H2O; Groves, 2006). The hydroxyl radical reacts with essentially all compounds in living organisms. The next reduction step leads to HO or H2O, the full reduced oxide (O2) state in which oxygen gains a saturated shell (neon shell). The reduction of oxygen by protein-bound reduced metals, flavins and some other reducing compounds is the cause for the uncontrolled generation of reactive oxygen species, the ‘‘toll’’ for aerobic life. Because they have devastating effects, they require immediate detoxification by antioxidant systems. Organisms apparently do not make use of photochemical activation to achieve controlled biochemical reactions of oxygen with organic compounds with oxygen; rather, accumulation of singlet oxygen is effectively counteracted by quenching (e.g., with carotenoids or vitamin E) to avoid damage. So far known, living organisms activate oxygen for controlled biochemical reactions always chemically by ‘‘dark’’ reactions. Similar as reduction of O2 as terminal electron acceptor occurs in a controlled manner in the oxidases (proteins that reduce oxygen with electrons) of electron transport chains, O2 reduction is also harnessed to generate reactive oxygen species for a controlled direct reaction with organic compounds in oxygenases. This can be more or less specific. Controlled generation of reactive oxygen species mostly takes place in iron-bound form, but sometimes also in copper-, flavin- or pterin-bound form.

3.2

Some Common Principles of Oxygenases

Oxygenases are highly diverse with respect to substrate specificity, mechanisms, active sites and phylogenetic relationships. Oxygenases are by no means used exclusively in the metabolism of hydrocarbons. They are also, and probably to a much larger extent, involved in a vast number of biosynthetic, detoxification and biodegradative reactions of many nonhydrocarbon (polar) compounds in all aerobic organisms, often to introduce polar groups at non-reactive (non-activated) carbon atoms or in a simpler way than via reaction sequences without oxygen. The usages of oxygenases for the activation of true hydrocarbons as notably unreactive compounds (> Table 1) thus represent special microbial adaptations of more general, wide-spread reaction principles. The hydroxylation of a polar compound may very much resemble that of a hydrocarbon if activation of the former occurs distantly from an activating group. For instance, omega-hydroxylation of a long-chain fatty acid (Coon, 2005) is like the hydroxylation of an n-alkane, even though a fatty acid may not ‘‘fit’’ into the substratebinding site of an alkane monooxygenase.

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

14

. Table 1 Overview of aerobic hydrocarbon activation in living organisms

Hydrocarbon(s) Methane

Short-chain non-methane alkanes (C2 to ca. C10)

Organisms

Enzyme type for activation

Only bacteria [Fe2]-Monooxygenase, soluble (sMMO) (aerobic methanotrophs) [Cu3]-Monooxygenase, particulate (pMMO) Bacteria

Long-chain non-methane Bacteria alkanes, (ca. > C10)

Remarks In few species of aerobic methanotrophs In essentially all species of aerobic methanotrophs

[Fe2]-Monooxygenase

[Heme]-Monooxygenase (P450-type) [Fe2]-Monooxygenase

In bacteria probably most common

[Flavin]-Monooxygenase

One case reported; mechanism unknown

Yeasts (mainly Candida sp.)

[Heme]-Monooxygenase (P450-type)

Yeasts may use alkanes as real growth substrate

Animals

[Heme]-Monooxygenase (P450-type)

By-reaction (‘‘detoxification’’) without further oxidation.

Alkenes

Bacteria

[Fe2]-Monooxygenase

Epoxide formation

Aromatic hydrocarbons

Bacteria

[Fe]-Diooxygenase

Very common for aromatic hydrocarbon utilization; formation of cis-hydrodiols

[Fe2]-Monooxygenase

Mono-hydroxylation of ring or alkyl side chain

[Flavin]-Monooxygenase

Epoxidation of styrene at the side chain

Some [Heme]-Monooxygenase filamentous (P450-type) fungi (e.g., Cunninghamella)

Mono-hydoxylation of ring, or formation of epoxides yielding trans-hydrodiols. Role as sole growth substrate uncertain

Mammals

Formation of epoxides; may yield transhydrodiols. By-reaction (‘‘detoxification’’) without further oxidation.

[Heme]-Monooxygenase (P450-type)

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Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

Oxygenases that activate hydrocarbons are always multi-component systems. A common feature is the electron transport in two subsequent one-electron steps to dioxygen to convert this into the oxidation state of a peroxide. From the peroxide state, a highly reactive oxygen species is generated by a disproportionation-like electronic rearrangement, without further net reduction by external electrons. The generated oxygen species then reacts with the saturated or unsaturated hydrocarbon3; the principle is depicted in > Fig. 2 for two important bacterial reactions, the activation of an alkane and of an aromatic hydrocarbon. Because NADH and NADPH as very common ‘‘electron’’-carrying coenzymes are restricted to transferring a hydride (H, equivalent to H+ + 2 e), generation of two single electrons requires a flavin (as prosthetic group of the reductase moiety of the oxygenase complex). The flavin accepts the hydride at the N5-position of the isoalloxazine ring and, by its ability to form a radical (semiquinone) state, releases single electrons subsequently to a redox-active iron center; this is often a Rieske-type Fe2S2 center, but other FexSx centers or a rubredoxin (containing single Fe coordinated by 4 cysteine; van Beilen et al., 2002) may be also involved. Via this iron center, the electrons reach stepwise the oxygenase component containing the O2-binding metal as the active site of carbon oxygenation. Electron transport to the active site may again involve an Fe2S2 center. An electron transport chain may be as follows (brackets indicate the protein): NADH ! H ! ½Flavin ! e þ e ! ½Fex Sx center ! e þ e ! ½metal center

O2

The metal center attains the appropriate (positive enough) redox potential for electron uptake upon binding of the hydrocarbon substrate to the enzyme. Also oxygen binding occurs after hydrocarbon binding and usually also after reduction of the metal center. There are quite different types of active sites in oxygenases. Furthermore, there is also variation of the electron transport components and subunit composition. A comprehensive classification of oxygenases with consideration of the different active sites and variation of all electron transport components and substrates (hydrocarbons, non-hydrocarbons) would be a tedious to almost impossible task. > Table 1 summarizes the different enzymatic modes for aerobic hydrocarbon activation in living organisms only in a rather general manner.

3.3

Heme-Iron Monooxygenases

Heme-iron monooxygenases are the most wide-spread and probably most diverse C–H-bond activating enzymes in living organisms (Denisov et al., 2005; Groves, 2006; Isin and Guengerich, 2007; Munro et al, 2007), even though they may not be the most frequently employed ones in bacterial activation of various hydrocarbons. There is a vast database of P450 protein structures and phylogenetic trees (http://www.ncbi.nlm.nih.gov/sites/entrez,‘‘structure’’; http://drnelson.utmem.edu/P450trees.html). They are also referred to as P450-type

3

Hence, the formal oxidation state of the carbon changes by +II. Assignment of formal oxidation states to the involved C–atoms before and after oxygenation may thus be used to check consistency of the formulated activation reaction. Examples: In terminal alkane oxygenation, the methyl group (CH3, III) is converted to a hydroxymethyl group (CH2OH, I). In an aromatic hydrocarbon, the (formally localized) ‘‘vinylen’’ (─CH=CH─, 2  I = II) can be dioxygenated yielding a hydrodiol (CHOHCHOH, 2  0 = 0) and a non-aromatic ring, or can be monooxygenated yielding an ‘‘enol’’ (CH=COH, I + I = 0) with maintenance of the aromatic ring. However, because the two electrons for O2 reduction are derived from an intermediate of hydrocarbon degradation, oxygenation totally consumes 4 electrons from the hydrocarbon substrate (> Fig. 2).

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

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. Figure 2 Principle of hydrocarbon oxygenation. O2 is always reduced to a bound peroxide state, [O22]; this directly yields the reactive oxygen species that attacks the hydrocarbon. (a) Reaction of monooxygenase (bifunctional oxygenase) with subsequent dehydrogenations by other enzymes. In some cases, the subsequent oxidations were partly attributed to by-reactions of the monooxygenase. (b) Reaction of dioxygenase with subsequent dehydrogenation (re-aromatization) by another enzyme.

hydroxylases (abbreviated CYP, followed by a number) because of their characteristic absorption band at 450 nm of the carbon monoxide complex. Of all oxygenases, heme-iron hydroxylases have been studied most intensely from a structural and mechanistic perspective, and their mechanism is the best-understood among oxygenase mechanisms. The bacterial enzyme P450cam, which hydroxylates the keto terpenoid, camphor at a non-activated methylene (– CH2–) group, is the longest-known of all hydroxylases (Hedegaard and Gunsalus, 1965, Schlichting et al., 2000). Several crystal structures of P450 enzymes have been elucidated (some examples from different domains of life: Cryle and Schlichting, 2008; Oku et al., 2004; Poulos et al., 1987; Smith et al., 2007). A consensus mechanism is depicted in > Fig. 3. The crucial step is the conversion of the O2-derived heme-iron-bound hydroperoxide to the highly reactive ferrylIV-species (FeIV=O) hosted in an oxidized porphyrin (lacking one p-electron). Formerly, this complex was usually regarded as a ferrylV-heme. P450 enzymes are the typical oxygenases in eukaryotes where they catalyze numerous specific or unspecific reactions mostly of non-hydrocarbons. Specific P450 enzymes are, for instance, involved in the synthesis of steroids or omega-hydroxylation of fatty acids (Coon, 2005). Unspecific P450 enzymes are involved in mammalian detoxification of various aliphatic and aromatic compounds (endogeneous compounds; contaminants from air, water and food; drugs) by increasing their hydrophilicity or reactivity. Nevertheless, activation may accidentally lead to a significantly increased toxicity, as in the case of the polycyclic aromatic hydrocarbon, benzo[a]pyrene that is converted to a carcinogen (Baird et al., 2005). The mammalian products of aromatic hydrocarbons are epoxides (arene oxides). These are either converted

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. Figure 3 Principle of monooxygenation of hydrocarbons by heme (P450-type) monooxygenases (hydroxylases). The porphyrin ring (Por) has donated an electron to the iron center, leaving a p–cation radical (Porþ ); the system is isoelectronic with Por–FeV=O. The favored mechanism is hydroxyl group formation via H-atom abstraction (R··H··O=FeIVPorþ ), resulting in an enzyme-bound alkyl radical (R ) and iron-coordinated hydroxyl radical (HO–FeIV/Por0), with subsequent recombination (rebound mechanism). For clarification of net charges, the sum of the charges and/or oxidation states of individual components of the complex are simply added. If the two carboxyl groups of protoporphyrin IX (Por) are regarded as undissociated, the iron-free form would be Por0. Heme formation by chelation of Fe2+, (which needs removal of 2 H+ from tetrapyrrole-nitrogen) leads to [Por2Fe2+]0. Binding of O2 followed by reduction with one ‘‘internal’’ electron from Fe2+ and one external electron leads to [Por2Fe3+(O–O)2], and protonation to [Por2Fe3+(O–OH)]0. Addition of another H+ and elimination of H2O yields [Por2Fe5+=O2]+. ‘‘Internal’’ donation of a p–electron from the porphyrin ring to the iron leads to [Por +/2Fe4+=O2]+. If the axial cysteine thiolate ligand is included, the complex is [RSPor +/2Fe4+=O2]0. Dissociation of the porphyrin carboxyl groups would yield [RSPor +/4Fe4+=O2]2.

to trans-dihydrodiols by epoxide hydrolases (> Fig. 4a), or rearranged by an NIH shift4 (intramolecular shift of a hydrogen by one position) to monohydroxyl arenes. However monohydroxyl arenes may be also formed without an expoxide as intermediate (> Fig. 4b;

4

Named after the National Institute of Health where this hydrogen shift was first detected (Guroff et al., 1967).

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

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. Figure 4 Oxygenation of aromatic hydrocarbons by P450 type oxygenases. The reaction is exemplified with benzene, C6H6 (mostly modified from Bathelt et al., 2008). (a) Formation of an epoxide with subsequent hydrolysis or intramolecular rearrangement (with NIH-shift). (b) Monohydroxylation of an aromatic hydrocarbon. Hydrogen atoms are only shown if they are relevant for the mechanism.

Bathelt et al., 2008). Even alkanes may be accidentally hydroxylated in mammals, most probably also by P450 enzymes (Crosbie et al., 1997; Frommer et al., 1972; Morohashi, 1982; Perbellini et al., 1980). In bacterial hydrocarbon activation, P450-type hydroxylases are often used for activation of alkanes of medium chain lengths (van Beilen et al., 2006; van Beilen and Funhoff, 2007). With respect to specificity, bacterial P450-type monooxygenases take an intermediate position. A given enzyme may accept a restricted number of similar (homologous) hydrocarbons. Yeasts which grow with alkanes also make use of P450-type monooxygenases (Ka¨ppeli, 1986; Scheller et al., 1996). The capability of P450-monooxygenases to activate aromatic compounds plays a role in fungi such as Cunninghamella elegans to initiate degradation (Cerniglia, 1992). As in mammals, oxygenation leads to an epoxide or aromatic mono-hydroxyl compound. However, fungal degradation of aromatic hydrocarbons is apparently incomplete (Bumpus, 1989; Wolter et al., 1997).

3.4

Non-Heme Di-Iron Monooxygenases

Non-heme di-iron monooxygenases are the most versatile and most frequently employed enzymes for aerobic hydrocarbon activation in bacteria. Their versatility surpasses that of

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P450 enzymes in hydrocarbon activation. There are soluble (Leahy et al., 2003; Lippard, 2005) and membrane-bound di-iron monooxygenases. The best-studied of the soluble di-iron enzymes is soluble methane monoxygenase (sMMO). It is present in some aerobic methanotrophic bacteria, whereas most aerobic methanotrophs possess the particulate tri-copper methane monooxygenase (see below; Murrell et al., 2000). Soluble methane monoxygenase has a very relaxed substrate specificity, reacting with numerous compounds such as alkanes or isoalkanes up to C8, methylsubstituted cylcopentane and cylcohexane, propylene, benzene, ethylbenzene, chloroethylenes, and halobenzenes (Burrows et al., 1984; Green and Dalton, 1989; Lipscomb, 1994). The crystal structure has been elucidated (Rosenzweig et al., 1993; Whittington and Lippard, 2001), and the mechanism has been studied (Han and Noodleman, 2008; Yoshizawa and Yumura, 2003; > Fig. 5). Phylogenetically related soluble di-iron monooxygenases epoxidize alkenes, or monohydroxylate toluene or o-xylene at the ring to yield p-cresol or o-xylene to 3,4-dimethylphenol (Fosdike et al., 2005; Leahy et al., 2003). The crystal structure of the latter enzyme type from Pseudomonas stutzeri has been elucidated (Sazinsky et al., 2004). The mechanism may include the formation of an epoxide, followed by a rearrangement via an NIH shift to yield the monohydroxylated compound. An extensively studied member of the membrane-bound di-iron monooxygenases is the alkane-monooxygenase AlkB from Pseudomonas putida (Bertrand et al., 2005). The membrane-bound methyl-hydroxylating toluene/xylene monooxygenase XylM (forming benzyl

. Figure 5 Monooxygenation by di-iron monooxygenase. The best studied enzyme is soluble methane monooxygenase. Of the ligands, only one characteristic bidentate carboxylate-group of a glutamate is shown. The reaction sequence has been simplified. Here, the rebound mechanism has been assumed. The FeIV–OH group is isoelectronic with FeIII– OH.

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

14

alcohol or methylbenzyl alcohols) encoded by the TOL plasmid in the same organism also involves a diiron center (Austin et al., 2003). Aerobic fatty acid desaturases in bacilli, cyanobacteria, yeasts and higher plants involve soluble enzymes with di-iron centers (Shanklin and Cahoon, 1998). They act upon acyl-ACP (carrier protein-bound fatty acids). The reaction takes place distant from any activating group (the thioester is thus not needed from a chemical point of view) and thus resembles an enyzmatic attack on an alkane.

3.5

Particulate Methane Monooxygenase (Particulate Di-Metal or Tri-Metal Monooxygenase)

Particulate methane monooxygenase (pMMO) is almost ubiquitous in aerobic methanotrophic bacteria (Murrell et al., 2000). The enzyme exhibits less substrate promiscuity than soluble methane monooxygenase, but alkanes up to C5 are also hydroxylated (Burrows et al., 1984); it also oxidizes ammonia to nitrite (Be´dard and Knowles, 1989). The enzyme has been crystallized and a tri-copper center was proposed to be involved in methane oxygenation (Balasubramanian and Rosenzweig, 2007; Chan and Yu, 2008; Liebermann and Rosenzweig, 2005; > Fig. 6). However, the metal composition is still a matter of dispute and may depend on the

. Figure 6 Assumed mechanistic principle of oxygenation by the tri-copper particulate methane monooxygenase, the most wide-spread methane activating enzyme in aerobic methanotrophs (modified and combined from Chan and Yu, 2008; Chen et al., 2007; Han and Noodleman, 2008). The presence of three copper ions (coordinated by histidine residues; not shown) in the native enzyme is still a matter of discussion, and the (insertion) mechanism is partly still speculative. Alternatively, a di-iron center was proposed for the reaction center (Martinho et al., 2007).

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bacterial species investigated (Hakemian et al., 2008); there is also spectroscopic evidence for a di-iron center, as in sMMO (Martinho et al., 2007). Particulate methane monooxygenase is evolutionarily related to the membrane-bound ammonia monooxygenase (AMO5) of ammonia-oxidizing bacteria (Be´dard and Knowles, 1989; Erwin et al., 2005; Zahn et al., 1996). AMO can oxidize methane to methanol, even though ammonia-oxidizing bacteria cannot grow with methane. Neither can methanotrophs that oxidize ammonia grow by this reaction.

3.6

Non-Heme Mono-Iron Oxygenases (Ring Hydoxylating Dioxygenases)

Activation of aromatic hydrocarbons by introduction of two hydroxyl functions to yield nonaromatic cis-hydrodiols is the most common activation mechanism in the degradation of aromatic hydrocarbons (See > Chapter 5 in this volume; Reinecke, 2001). Numerous ringhydroxylating dioxygenases have been described and classified (See chapter by Pe´rez-Pantoja et al. in this volume; Kweon, 2008). Enzymes performing dihydroxylation of rings contain a single-iron center coordinated by amino acid residues. The crystal structure of naphthalene dioxygenase has been determined (Karlsson et al., 2003). Details of the reaction mechanism are still a matter of discussion (Bugg, 2003). A favored mechanism is presented in > Fig. 7. Dioxygenases may also possess monooxygenase activity towards some compounds; for instance naphthalene dioxygenase can introduce a single hydroxyl group into the hydrocarbons indene (fused aromatic six- and five-membered rings) and ethylbenzene, in the latter at carbon-1 of the side chain (Resnick et al., 1996). Ring-activating dioxygenases have nothing in common with ring-cleaving dioxygenases. The latter act upon the aromatic ortho-dihydroxy compounds formed by activation. The ring-cleaving enzymes do not involve an electron transport from an external donor to O2, and they do not activate hydrocarbons (See > Chapter 18, Vol. 2, Part 4; > Chapter 4, Vol. 2, Part 2; Bugg, 2003).

3.7

Flavin-Containing Monooxygenases, and Monooxygenases with Unknown Mechanism

Monooxygenases without metal but with a flavin in the reaction center are involved in hydroxlyations in bacteria and eukaryotes. The oxygenating species is the flavin-4a-hydroperoxide (> Fig. 8). Flavin-containing hydroxylases usually hydroxylate organic compounds at carbon atoms activated by polar groups, for instance phenolic compounds, or they oxygenate organic heteroatoms such as nitrogen and sulfur (See > Chapter 18; van Berkel et al., 2006). Reports about oxygenation of true hydrocarbons by flavin-containing monooxygenases are scarce. A case studied in detail is the epoxidation of the synthetic hydrocarbon styrene (C6H5–CH=CH2) at the relatively reactive side chain (> Fig. 8) by a two-component flavin-monooxygenase belonging to so so-called subclass E out of six subclasses (Beltrametti et al., 1997). 5

Abbreviation AMO must not be confused with the same one sometimes used for alkene monooxygenase.

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. Figure 7 One of the suggested mechanisms in dioxygenation of aromatic hydrocarbons by mono-iron dioxygenase (combined and modified from Bugg, 2003; Chakrabarty et al., 2007; Kovaleva et al., 2007). The iron is coordinated by two His and one bidentate Asp. Other suggested alternatives are a radical mechanism (AR + O=FeV–OH !  AR–O–FeIV–OH ! HO–AR–O–FeIII) or formation of an arene expoxide as intermediate.

A surprising finding was the recovery of a flavin-monooxygenase from an alkanedegrading bacterium. From Geobacillus thermodenitrificans, a gene for a protein (mainly found in the extracellular fraction) was expressed in E. coli and yielded a protein converting longchain alkanes (C15–C36) to 1-alkanols, initially without obvious coenzyme requirement (Feng et al., 2008). Subsequently, the crystal structure was determined and the protein, a member of the bacterial luciferase family, was found to bind FMN. It was proposed to hydroxylate long-chain alkanes with FMN-4a-hydroperoxide (Li et al., 2008). The unusual reaction awaits elucidation. In an Acinetobacter strain able to degrade various alkanes, transposon mutagenesis revealed various genes needed for alkane metabolism (Throne-Holst et al., 2007). A gene was found to be required for the metabolism of C32 and C36 alkanes; there was a close relationship to a gene from another Acinetobacter strain able to utilize polar aromatic compounds. The gene was assumed to encode a putative flavin-binding monooxygenase. A dimeric protein purified from an Acinetobacter strain was reported to catalyze O2-dependent consumption of n-alkanes from C10 to C30 (Maeng et al., 1996). Whereas stoichiometric oxygen consumption and product formation during alkane consumption could not be measured, formation of non-stoichiometric traces of terminal alkyl hydroperoxides (R–CH2–OOH) was observed. The stoichiometry and mechanism of the reaction need

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. Figure 8 Oxygenation of the (synthetic) hydrocarbon, styrene, by flavin-monooxygenase (hypothesized in this article according to information by Ballou et al., 2005; van Berkel et al., 2006), yielding an epoxide. Flavin-monooxygenases, which perform numerous hydroxylations of nonhydrocarbons, are scarcely involved in hydrocarbon activation. The activation of styrene, a rather reactive hydrocarbon, is the best studied example. Reduced flavin formed by H transfer from NADH transfers an electron to O2 yielding flavin semiquinone and superoxide (FADH + O2 ! FADH + O2 , not shown). Addition of the superoxide, which requires a spin-inversion, and protonation leads to the 4a-hydroperoxide (FADH–O–OH). The hydroperoxide reacts as a nucleophile with the p-bond of the side-chain. The formed epoxystyrene is converted by another enzyme, an isomerase, to phenylacetaldehyde (Beltrametti et al., 1997). The mechanism for an involvement of a flavin-4a-hydroperoxide in the hydroxylation of alkanes as very stable compounds with only s-bonds is matter of discussion.

further investigation. An electron donor such as NAD(P)H was not needed. The protein contained FAD and was stimulated by copper. Formation of an aldehyde as subsequent product was discussed.

4

Anaerobic Activation of Hydrocarbons

The low reactivity of many hydrocarbons and the need for highly reactive, O2-derived coreactants to achieve metabolic activation have been arguments in favor of the former view that hydrocarbons are biologically inert in the absence of oxygen. However, observations in natural microbial populations and subsequently in many cultures provided a growing body of evidence for a strictly anaerobic degradation of saturated and unsaturated hydrocarbons (for review see Spormann and Widdel, 2001; Widdel and Rabus, 2001; Widdel et al., 2007). Anaerobic hydrocarbon degraders grow significantly slower than their aerobic counterparts (doubling times of a day to weeks vs. some hours); this may be one reason for the relatively late discovery and

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

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cultivation of anaerobic hydrocarbon degraders. Whereas aerobic activation always involves activated, oxidizing forms of oxygen, anaerobic activation mechanisms involve substantially different types of co-reactants to introduce polar groups. These may be polar carbon compounds, oxygen at the oxidation state of water (fully reduced oxygen) or even a transition metal forming a transient carbon-metal bond. The principal types of enzymes are listed in > Table 2.

4.1

Anaerobic Activation of Methane

The anaerobic oxidation of methane (AOM) with sulfate is a process that is wide-spread in marine sediments (See > Chapter 7 in this volume). Because it controls the emission of methane as a greenhouse gas, AOM is of global significance. AOM is mediated by associations of archaea (related groups of anaerobic methanotrophs, ANME) and bacteria related to sulfatereducing bacteria of the Deltaproteobacteria. These associations are usually interpreted as a

. Table 2 Overview of anaerobic hydrocarbon activation s in living organisms (for the alkyne, acetylene, see Seiffert et al., 2007). Anaerobic hydrocarbon activation has not been observed in eukaryotes

Hydrocarbon(s)

Organisms

Enzyme type for activation

Remarks

Archaea (assumed syntrophism with SO42-reducing bacteria)

Ni-porphinoid enzyme (closely related to CH3– coenzyme M reductase)

Assumed reversal of the final step in methanogenesis

Bacteria (NO3reducing)

Enzyme unknown

Different from the final step in methanogenesis

Bacteria

Glycyl radical enzyme

Leads to a substituted succinate

Unknown enzyme

Apparent activation at carbon-3

Bacteria

Unknown enzyme

Introduction of an HOgroup

Benzene, naphthalene, phenanthrene

Bacteria

Unknown enzymes

Activation results in addition of a COO-group (methylation also discussed)

Methylbenzenes

Bacteria

Glycyl radical enzyme

Leads to a substituted succinate

Ethylbenzene

Bacteria, SO42reducing

Glycyl radical enzyme

Leads to a substituted succinate

Mo-cofactor-containing dehydrogenase

Leads to (S)-1phenylethanol or (S)-1phenylpropanol

Methane

Non-methane alkanes

Alkenes Aromatic hydrocarbons

Ethylbenzene, Bacteria, NO3n-propylbenzene reducing

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. Figure 9 Hypothesized anaerobic activation of methane by a thiyl radical vs. activation by a high-valent nickel (drawn according to discussions by Kru¨ger et al., 2003; Shima and Thauer, 2005; Thauer and Shima, 2008) (a) In the initially suggested mechanism, the heterodisulfide of two coenzymes (CoB–S–S–CoM) is reduced by cofactor F430-hosted NiI as a strong reductant. This leads to a NiII-coordinated thiolate and a thiyl radical (CoB–S ) that attacks the methane. The methyl radical ( CH3) combines with the thiolate to form methyl-coenzyme M and to regenerate the NiI-state. (b) The presently favored mechanism begins with the reduction of the heterodisulfide to yield the NiIII-state as a strong electrophile. This attacks the methane to combine with a methyl carbanion (CH3; not depicted), viz. yielding a metal-organic compound (NiIII–CH3). Then the methyl group is transferred as a methyl carbenium ion equivalent (CH3+; not depicted) to the coenzyme M thiolate to form methyl-coenzyme.

syntrophism in which the archaeal partner activates the methane and the bacterial partner scavenges a methane-derived electron donor from the archaeon to reduce sulfate. The net reaction is according to CH4 þ SO42 þ Hþ ! HCO3 þ H2S þ H2O. The close relationship of the involved archaea to methanogenic archaea as well as high similarities between ANME group-derived proteins (> Fig. 10) and encoding genes to those of methanogenic archaea led to the conclusion that AOM is essentially a reverse methanogenesis. Accordingly, the methane-activating enzyme is thought to be a reverse methyl-coenzyme M reductase, a nickelporphinoid enzyme catalyzing the terminal step in methanogenesis. Hence, also the mechanisms of methane oxidation and methane formation are expected to be essentially similar, the main difference being the direction of the reaction. The presently discussed mechanisms (> Fig. 9) are (a) an attack by a thiyl radical (the coenzyme B radical, CoB–S ) on methane (Kru¨ger et al., 2003), or (b) a reaction of cofactor F430 in its Ni(III) state as a strong electrophile to form a Ni(III)-methyl compound (Shima and Thauer, 2005; Thauer and Shima, 2008). In the latter mechanism, a Ni(III)-hydride may play a role (Harmer et al., 2008).

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. Figure 10 Relationships of the assumed anaerobic methane-activating enzyme (reverse methyl-coenzyme M reductases, rMcr) in anaerobic methanotrophs (bold) to regular methyl-coenzyme M reductase (Mcr) in methanogenic archaea (drawn according to Hallam et al., 2003; Kru¨ger et al., 2003; Lo¨sekann et al., 2007; A. Meyerdierks, personal communication). The tree is based on genededuced amino acid sequences of the large subunits (rMcrA, McrA). The protein from ANME-1 was purified from naturally highly enriched methanotrophic mats (Kru¨ger et al., 2003), so as to confirm the underlying gene. Assignment of the other gene-derived proteins to methanotrophs (rather than to methanogens) was based on the natural abundance of ANME cells in a habitat, or on base composition statistics.

A different mode of methane activation may take place in a highly enriched methaneoxidizing denitrifying culture (Raghoebarsing et al., 2006). Whereas the early enrichment still contained archaea, subcultures were essentially dominated by a bacterial phylotype, and involvement of a reverse methyl-coenzyme M reductase was excluded (Ettwig et al., 2008). The mechanism is still unknown. A radical-catalyzed addition to fumarate to yield methylsuccinate, analogous to the activation of non-methane alkanes (see next section), is speculative.

4.2

Glycyl Radical Enzymes for Anaerobic Activation of Non-Methane Hydrocarbons

The finding of benzylsuccinate as a metabolite in toluene-degrading anaerobic cultures (Beller et al., 1992; Evans et al., 1991) and subsequent enzymatic measurements (Beller and Spormann, 1997; Biegert et al., 1996) led to the discovery of a frequently occurring anaerobic activation mechanism for alkyl-substituted aromatic hydrocarbons, especially those with methyl groups. The activation occurs at the sp3-carbon adjacent to the aromatic ring. The

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reaction (> Fig. 11) has been studied in detail with the toluene-activating benzylsuccinate synthase from denitrifying Betaproteobacteria (See > Chapter 15 in this volume; Heider, 2007). According to metabolite studies, the aromatic hydrocarbon ethylbenzene in sulfate-reducing bacteria is also activated via a radical-catalyzed addition to fumarate, yielding (1-phenylethyl)succinate (Elshahed et al., 2001; Kniemeyer et al., 2003). Ethylbenzene activation in denitrifiers occurs in a different manner (see next section). An essentially analogous reaction principle (> Fig. 11) is found in the anaerobic activation of many alkanes with three or more carbon atoms. They are usually activated at carbon2 yielding (1-methylalkyl)succinates. However, in the case of propane, activation at carbon-1 has been also observed, presumably in a by-reaction (Kniemeyer et al., 2007). Long-chain alkanes may also react at carbon-3 in a by-reaction (Rabus et al., 2001). Anaerobic ethane utilization (coupled to sulfate reduction) is an extremely slow process (Kniemeyer et al., 2007), and neither the involved organisms nor a potential intermediate have been identified. According to analyzed genes, the enzymes or candidate enzymes for anaerobic activation of alkyl-substituted aromatic hydrocarbons are related to each other (> Fig. 12).

4.3

Ethylbenzene Dehydrogenase, a Molybdo-Enzyme

Ethylbenzene in denitrifying bacteria is activated by dehydrogenation (anaerobic hydroxylation) to 1-phenylethanol (See > Chapter 15 in this volume; Johnson et al., 2001; Kniemeyer and Heider, 2001) rather than by addition to fumarate, as in sulfate-reducing bacteria (see above). The three-dimensional structure of the enzyme, ethylbenzene dehydrogenase, has been

. Figure 11 Principles of the anaerobic activation of alkanes or methylaryl hydrocarbons by addition to fumarate catalyzed by a glycyl radical enzyme. The glycyl radical is generated by reductive cleavage of S-adenosylmethionine into methionine and the adenosyl radical. The latter generates a glycyl radical (enzyme activation) and is converted to 50 -desoxyadenosine. The glycyl radical subsequently generates a thiyl radical that attacks the hydrocarbon in an indefinite number of cycles.

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

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. Figure 12 Relationships of anaerobic hydrocarbon-activating enzymes (bold) to other glycyl radical enzymes. (Modified from Musat et al., 2008).

resolved and a detailed mechanism has been suggested (See > Chapter 15 in this volume; Szaleniec et al., 2007; > Fig. 13). It belongs to the dimethylsulfoxide (DMSO) reductase family of molybdoenzymes (Bender et al., 2005); so far, no other hydrocarbon-activating enzymes are known in this family. In the mechanism, a Mo(VI) species withdraws a hydride. Such a unique dehydrogenation of a hydrocarbon is obviously possible by a high stabilization of the carbenium ion and an electron acceptor with high enough redox potential as achievable in the metabolism of a denitrifier.

4.4

Unknown and Hypothesized Reactions

Patterns of cellular fatty acids observed in the alkane-degrading sulfate-reducing strain Hxd3 (Aeckersberg et al., 1998; So et al., 2003) were not in accordance with a radical-catalyzed addition of carbon-2 to fumarate and the subsequent degradation involving carbon skeleton rearrangement and decarboxylation (Rabus et al., 2001; Wilkes et al., 2002). Based on labeling studies, addition of a carboxyl function at carbon-3 has been proposed as the initial enzymatic reaction (So et al., 2003). In the anaerobic oxidation of unsubstituted aromatic hydrocarbons such as benzene, naphthalene or phenanthrene, carboxylation (Caldwell and Suflita, 2000) has been most frequently discussed as the activation mechanisms. These suggestions were based on labeling studies and the identification of aromatic hydrocarbon-derived carboxylic acids with CO2derived carboxyl groups. In addition, naphthalene methylation followed by addition to fumarate was suggested as an alternative mechanism in particular cultures (Coates et al., 2002; Safinowski and Meckenstock, 2006; Ulrich et al., 2005). However, in marine cultures of benzene- or naphthalene-degrading sulfate-reducing bacteria, methylation was essentially excluded (Musat and Widdel, 2008; Musat et al., 2008). In the anaerobic degradation of alkenes, addition of water to the double bond has been hypothesized, in the case of 1-alkenes with anti-Markovnikov orientation (R–CH = CH2 + H2O ! R–CH2–CH2OH; Schink ,1985). Water addition would be particularly favored by involvement of a tertiary carbon atom, as occurring with monoterpenes (See > Chapter 12 this volume). In principle, also anaerobic hydroxylation at the carbon atom next to the

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. Figure 13 Proposed principle of the anaerobic hydroxylation (dehydrogenation) of ethylbenzene by ethylbenzene dehydrogenase (simplified and modified from Szaleniec et al., 2007; for additional information see also chapter by Boll and Heider, this volume). The reaction is unique in that there is enough stabilization of the carbenium cation to be formed oxidatively by hydride abstraction in an anaerobic metabolism.

double bond (the allylic carbon) appears possible (–CH2–CH=CH– + H2O ! –CHOH– CH=CH– + 2 [H]), analogous to ethylbenzene dehydrogenation at the benzyl carbon atom.

5

Research Needs

In aerobic oxygenation reactions, a broader understanding of the energetic prerequisites to achieve hydrocarbon activation by an oxygen species appears desirable. Activation of a hydrocarbon requires a ‘‘harsh’’ reactant in the active site. Why is the energetic state (‘‘reactivity’’) of oxygen in one oxygenase ‘‘high’’ enough to activate an non-methane alkane or even methane, whereas in another case the oxygen can only activate more reactive carbon atoms next to functional groups? Furthermore, the factors that determine substrate specificity of oxygenases are of much interest. Are conventional sterical ‘‘sustrate-fit’’ models sufficient to explain why some oxygenases have a strikingly broad substrate spectrum, for instance di-iron methane monooxygenase or mammalian detoxifying P450 enzymes, whereas some monooxygenases accept only a few alkanes (van Beilen and Funhoff, 2007). Combined effects of the energetic state of the oxygen and steric factors in substrate binding may play a role and thus require deeper understanding. Furthermore, one may ask whether an oxygenase that hydroxylates only organic compounds with activated carbon atoms (polar organic compounds) can be genetically engineered so as to achieve activity towards a hydrocarbon.

Diversity and Common Principles in Enzymatic Activation of Hydrocarbons

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A fatty acid hydroxylase (hydroxylating at subterminal carbon) could be converted to an alkane hydroxylase (Fasan et al., 2007); the reactions per se resemble each other. This leads directly into the field of oxygenase engineering, which is of significant biotechnological interest for fine chemical production from hydrocarbons and non-hydrocarbons (Boyd et al., 2001; Ullrich and Hofrichter, 2007; Urlacher and Schmid, 2006, van Beilen and Funhoff, 2005). Such experiments may also shed light on the natural evolution of hydrocarbon activating oxygenases that may have evolved from enzymes activating more reactive polar compounds. Also health-related aspects of human (or mammalian) oxygenases that react with hydrocarbons are of significant interest. Some aromatic compounds are procarcinogens because one of their subsequent oxygenation products can transform normal cells into cancer cells. In the case of benzo[a]pyrene, the carcinogenic effect is attributed to the benzopyrene hydrodiol epoxide that binds covalently to guanine in DNA (Baird et al., 2005). However, in the case of benzene, another potential procarcinogen, the epoxide may not be the cell-transforming agent (Golding and Watson, 1999; Powley and Carlson, 2000). More studies of reactions between hydrocarbon activation products and DNA are needed to understand why some aromatic hydrocarbons are potential carcinogens whereas several others are not. The anaerobic, oxygen-independent hydrocarbon activation is a much younger research area than aerobic activation, and anaerobic mechanisms are far less understood than aerobic mechanisms. Principles in anaerobic hydrocarbon activation are apparently more diverse than in aerobic activation; in anaerobes, the hydrocarbons may undergo combination with other carbon compounds, hydroxylation via dehydrogenation, or addition of a transition metal. The activation of unsubstituted aromatic hydrocarbons is poorly understood. Furthermore, there might be alternative anaerobic activation mechanisms of alkanes not involving a radicalcatalyzed addition to fumarate. As with oxygenases, an understanding of the energetic state of the active site component that achieves hydrocarbon activation (such as C–H cleavage) is also of significant interest in anaerobic hydrocarbon activation. Furthermore, the observation that bacteria degrading hydrocarbons anaerobically have a very narrow substrate range (with respect to hydrocarbons), presumably due to high specificity of activating enzymes, is not understood. And, again as in the case of aerobes, insights into the evolution of the anaerobic hydrocarbonactivating enzymes from ancestors that activate other substrates are of basic interest.

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15 Anaerobic Degradation of Hydrocarbons: Mechanisms of C–H-Bond Activation in the Absence of Oxygen M. Boll1 . J. Heider2 Insitute of Biochemistry, University of Leipzig, Leipzig, Germany [email protected] 2 Laboratory for Microbiology, University of Marburg, Germany [email protected] 1

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Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1012

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C–H-Bond Activation Energies and Catalytic Mechanism . . . . . . . . . . . . . . . . . . . . . . . 1013

3 3.1 3.2 3.3 3.4

Mechanisms of C–H-Bond Activation in Anaerobic Hydrocarbon Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1013 Anaerobic Hydroxylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1013 Addition of Fumarate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1016 Reverse Methanogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1017 Methylation and Carboxylation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1019

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1021

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_71, # Springer-Verlag Berlin Heidelberg, 2010

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Anaerobic Degradation of Hydrocarbons

Abstract: Hydrocarbons are highly abundant in nature and are formed either via geochemical or biological processes. Their high C–H bond dissociation energies are responsible for low chemical reactivities. Due to the toxicity of many hydrocarbons, their biological degradation is of environmental concern. In the presence of oxygen, the C–H-bond is activated by oxygenases involving enzyme-bound reactive oxygen species in exergonic reactions. In contrast, anaerobic hydrocarbon-degrading bacteria use a number of alternative enzymatic reactions for the mechanistically sophisticated C–H-bond activation. Some of these reactions are only known from anaerobic hydrocarbon degradation pathways, and some follow unprecedented biochemical mechanisms. The known oxygen-independent C–H-activation reactions comprise: (1) hydroxylation with water by molybdenum cofactor containing enzymes, (2) addition to fumarate by glycyl-radical enzymes, (3) reverse methanogenesis involving variants of methyl-coenzyme M reductase, (4) methylation, and (5) carboxylation catalyzed by yetuncharacterized enzymes. The available knowledge about these enzymes varies greatly: the ethylbenzene hydroxylating molybdenum enzyme has been characterized structurally and functionally, whereas even the mode of initial activation is at issue in case of benzene degradation (methylation vs. carboxylation).

1

Introduction

The enzymatic reactions involved in the aerobic degradation of hydrocarbons by bacteria or fungi have extensively been studied for several decades. Under oxic conditions, hydrocarbon metabolism is always initiated by mono- or dioxygenase reactions. The dioxygen molecule is usually activated by these enzymes to a highly reactive metal-bound oxygen species that enables the terminal or subterminal hydroxylation of aliphatic alkane chains or the monoor dihydroxylation of aromatic rings (Harayama et al., 1992; McLeod and Eltis, 2008). For this reason, the capability to fully degrade hydrocarbons has long been considered as an exclusive feature of aerobic microorganisms. However, it has meanwhile been established that anaerobic degradation of hydrocarbons also plays an important role in nature. For example, contaminated marine sediments or aquifers represent typical natural environments where oxygen as terminal electron acceptor is quickly depleted and hydrocarbons have to be degraded anaerobically. In the last two decades, the number of known anaerobic hydrocarbon-degrading bacteria has been increased continuously. In recent years, the first available genomes of hydrocarbon degrading anaerobes enabled easier access to the genes and enzymes involved in the degradation pathways. Studying anaerobic hydrocarbon metabolism has revealed unprecedented enzymatic reactions involved in C–H-bond activation, which are already well understood in a few cases (e.g., ethylbenzene hydroxylation), but still are at issue in others (e.g., anaerobic benzene degradation). A number of reviews focusing on different aspects of anaerobic hydrocarbon metabolism have been published in recent years (Boll et al., 2002; Chakraborty and Coates, 2004; Heider, 2007; Heider and Rabus, 2008; Thauer and Shima, 2008; Widdel and Rabus, 2001). This review gives an overview of the enzyme reactions involved in the initial C–H bond activation in anaerobic hydrocarbon degradation pathways. Some general considerations relating activation energies of C–H-bonds of different hydrocarbons to the types of individual enzymatic reactions involved are suggested, and the biochemical principles of characterized and proposed mechanisms of these enzymes are presented. With the exception of p-cresol, this review focuses mainly on hydrocarbons carrying no heteroatoms.

Anaerobic Degradation of Hydrocarbons

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15

C–H-Bond Activation Energies and Catalytic Mechanism

In > Table 1, the C–H-bond dissociation energies of some selected aliphatic and aromatic hydrocarbons are listed. The range spans over 100 kJ mol
1, which makes it obvious that the enzymes catalyzing initial hydrocarbon activation must use different mechanisms, depending on the amount of energy required for C–H-bond cleavage. The C–H bond dissociation energies of benzene and methane are highest among all hydrocarbons (>430 kJ mol
1). In aerobic organisms this high energetic barrier is overcome by using highly reactive oxygen species. In case of monooxygenases, a bound dioxygen molecule is partially reduced to water and an enzyme-bound oxo-ferryl species, which can be considered as an equivalent of hydroxyl radical held under control in the active site (Lieberman and Rosenzweig, 2004; Thauer and Shima, 2008). The formation of a free hydroxyl radical from H2O involves 497 kJ mol
1, which is sufficient to cleave any C–Hbond in an exergonic reaction. However, it is hard to reconcile the high energies of C–H-bond dissociation of benzene and methane with the observed widely distributed occurrence of their oxidation in anoxic environments in nature. As anaerobic bacteria cannot generate reactive oxygen species, alternative mechanisms are realized for benzene and methane activation. Very likely, these involve alternative enzyme radical species and probably couple exergonic to endergonic partial reactions (Thauer and Shima, 2008). Compounds with lower C–H bond dissociation energies in the range of 355–423 kJ mol
1 (ethylbenzene, toluene, aliphatic alkanes with chain lengths 2) up to possibly 439 kJ mol
1 (methane) are predominantly activated by enzymatic addition to a fumarate cosubstrate. Finally, compounds with lower C–H-bond dissociation energies (355 kJ mol
1) can also be degraded by anaerobic hydroxylation with water (e.g., ethylbenzene, n-propylbenzene, p-cresol). Such hydroxylations produce electron equivalents according to the following equation: R-H + H2O ! R-OH + 2[H]. Because of the relatively high standard potential of the reaction (e.g., E0 [1-phenylethanol/ethylbenzene] = + 30 mV), these reactions seem only possible under anaerobic respiratory conditions with electron acceptors of even higher standard potentials. This assumption is evident from the anaerobic degradation pathways of p-cresol and ethylbenzene, which proceed via methyl group hydroxylation in denitrifying bacteria (E0 of terminal electron acceptors + 430 mV), (Hopper, 1976; Johnson et al., 2001; Kniemeyer and Heider, 2001; McIntire et al., 1985), but via addition to fumarate in sulfatereducing bacteria (E0 of terminal electron acceptors Fig. 1).

3

Mechanisms of C–H-Bond Activation in Anaerobic Hydrocarbon Degradation

3.1

Anaerobic Hydroxylation

The hydroxylation of the methyl-group of p-cresol (4-methylphenol) by water has been demonstrated in aerobic, denitrifying bacteria and Fe(III)-reducing bacteria (Peters et al., 2007) and many details about the structure–function relationship of the corresponding p-cresol methyl hydroxylase have been obtained (Cunane et al., 2005, 2000; Efimov et al., 2004). This enzyme is a flavocytochrome abstracting a hydride from the methyl group and yielding a relatively stable methide quinone cation intermediate. Similar anaerobic

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. Table 1 C–H bond dissociation energies of hydrocarbons and type of reactions/enzymes/cofactors involved in hydrocarbon activation in anaerobic bacteria

Compound

Dissociation energy (kJ mol
1) 473

Mechanism of C–H-cleavage

Enzyme

Cofactors

Carboxylation?

Unknown

Unknown

Methylation?

Unknown

Unknown

Reverse methanogenesis

Methyl-CoM reductase

F430/Ni

Addition to fumarate?

Glycyl radical enzyme?

Glycyl radical?

423

Addition to fumarate

Glycyl radical enzyme?

Glycyl-radical/ FeS?

405

Addition to fumarate

Glycyl radical enzyme?

Glycyl-radical/ FeS?

376

Addition to fumarate

Benzylsuccinate synthase

Glycyl-radical/ FeS

355

Addition to fumarate

Glycyl radical enzyme

Glycyl-radical/ FeS?

439

335

Hydroxylation by Ethylbenzene water dehydrogenase

Mo-Cofactor FeS Heme b

Addition to fumarate

Glycyl-radical/ FeS?

Glycyl radical enzyme

Hydroxylation by p-Cresol FAD Heme c water methylhydroxylase Reactions/cofactors with questions marks have not been identified so far. Values for C–H-bond dissociation energies were taken from Thauer and Shima (2008)

hydroxylation reactions were initially suggested for nonphenolic alkylated aromatics. However, a hydride cannot be easily extracted from toluene as no quinone intermediate can be formed. This can be derived from the much higher C–H-bond dissociation energy of toluene compared with that of the phenolic analog p-cresol (> Table 1). Therefore, different types of enzymes are required for the activation of nonphenolic alkylbenzenes (see 2.2). The C–H-bond dissociation energy at C-2 of ethylbenzene lies between the values of p-cresol and toluene. It has been demonstrated that ethylbenzene is initially attacked by hydroxylation with water yielding (S)-phenylethanol and two electron equivalents. The corresponding ethylbenzene dehydrogenases have been studied in two different denitrifying

Anaerobic Degradation of Hydrocarbons

15

. Figure 1 Initial activation reactions involved in anaerobic degradation of toluene (a), ethylbenzene (b) and p-cresol (c) in different anaerobic, hydrocarbon degrading bacteria.

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Anaerobic Degradation of Hydrocarbons

bacteria and the enzyme from Aromatoleum aromaticum meanwhile represents the best-characterized enzyme in anaerobic hydrocarbon degradation (Johnson and Spormann, 1999; Johnson et al., 2001; Kniemeyer et al., 2003). Ethylbenzene dehydrogenase is a molybdenum-cofactor (MoCo) containing enzyme of the dimethylsulfoxide reductase family. The structure of the enzyme from Aromatoleum aromaticum (formerly Azoarcus EbN 1) has been solved at 1.88 A˚ (Kloer et al., 2006). It contains three subunits: the a-subunit carries a molybdenum-bis-molybdopterin guanine dinucleotide cofactor and a [4Fe-4S] cluster, the b-subunit carries four further FeS-cluster, and the g-subunit represents a b-type cytochrome. Based on structural and kinetic data, a reaction mechanism was proposed that is initiated by hydride transfer from C-2 of the substrate to a Mo(VI) = O species of the cofactor (Kloer et al., 2006; Szaleniec et al., 2007). This reaction generates a substrate carbenium cation and a Mo(IV)–OH species in the transition state. After stereospecific hydroxylation of the cation with the hydroxide coordinated to the Mo-(IV), the product (S)-phenylethanol is released by hydrolytic cleavage. Finally, the Mo(VI) species is regenerated by two single-electron transfer steps via the FeS-clusters to the cytochrome b-subunit (> Fig. 2). Very recently, a similar reaction has been discovered in the anaerobic bacterial degradation of cholesterol in the denitrifying Sterolibacterium denitrificans. The tertiary carbon atom of the alkyl side chain from the intermediate cholest-1,4-diene-3-one is anaerobically hydroxylated by water, and the involvement of a molybdenum cofactor containing hydroxylase has been proposed (Chiang et al., 2007).

3.2

Addition of Fumarate

The discovery of benzylsuccinate as the initial intermediate of anaerobic toluene degradation (Beller et al., 1992; Evans et al., 1992; Biegert et al., 1996) led to a novel type of biochemical reaction, which often appears to be involved in anaerobic C–H-bond-activation: the reaction of alkyl chains with a fumarate cosubstrate yielding an alkylsuccinate adduct. Evidence for such a reaction has been demonstrated for the initial activations of toluene, xylenes, ethylbenzene, p-cresol, methylnaphthalene, cyclohexane, n-hexane, and short chain alkanes; some selected

. Figure 2 Proposal for reaction mechanism of ethylbenzene dehydrogenase.

Anaerobic Degradation of Hydrocarbons

15

references for original descriptions and reviews: (Biegert et al., 1996; Heider, 2007; Kniemeyer et al., 2003, 2007; Morasch et al., 2004; Mu¨ller et al., 2001; Rabus et al., 2001; Safinowski and Meckenstock, 2004; Selmer et al., 2005; Wilkes et al., 2002). In addition, a similar reaction has been discussed for anaerobic methane oxidation coupled to denitrification (Thauer and Shima, 2008). In the following the principles of C–H-bond activation by addition to fumarate are presented for benzylsuccinate synthase (BSS), the prototype of this class of enzymes. BSS has initially been isolated and characterized from the denitrifying bacterium Thauera aromatica (Leuthner et al., 1998). It catalyzes the first step in anaerobic toluene catabolism, the stereospecific syn-addition of toluene to fumarate yielding (R)-benzylsuccinate (Biegert et al., 1996; Qiao and Marsh, 2005). It belongs to the class of glycyl radical enzymes comprising pyruvate formate lyase or anaerobic ribonucleotide reductase (Selmer et al., 2005). All members of this class are extremely oxygen labile due to the presence of a glycyl radical in the peptide chain, which can be detected by its typical electron paramagnetic resonance spectrum (Krieger et al., 2001; Duboc-Toia et al., 2003; Verfu¨rth et al., 2004). Exposure to oxygen results in the irreversible peptide chain cleavage at the position of the radical species. The glycyl-radical is not considered to be directly involved in catalysis; it rather represents a relatively stable form of an enzyme radical that initiates the catalytic cycle by abstracting a proton from a cysteine residue forming a much more reactive thiyl ‘‘working’’ radical (Boll et al., 2002; Himo, 2005) (> Fig. 3). BSS from T. aromatica is composed of three subunits: a large subunit carrying a stable glycyl radical and two smaller subunits carrying FeS clusters. BSS catalysis has been proposed to be initiated by hydrogen atom abstraction from the methyl group of toluene yielding a benzyl radical. The benzyl radical then adds to fumarate yielding a benzylsuccinyl radical, which reabstracts a hydrogen atom from the enzyme (most possibly the conserved cysteine) yielding benzylsuccinate and the regenerated glycyl radical (> Fig. 3). The energetics of the benzylsuccinate synthase reaction are puzzling. The formation of a stable s–bond at the expense of the weaker C–C p–bond of fumarate is considered to be clearly exergonic (DG0 =
56 kJ mol
1), although some evidence for a very slow backward reaction has been obtained (Li and Marsh, 2006). On the other hand, formation of the glycyl radical by bond dissociation requires 350 kJ mol
l. This energy appears to be too low to explain the formation of the catalytically relevant thiyl radical (365 kJ mol
1) and the benzyl substrate radical (376 kJ mol
1). Considering that similar glycyl radical enzymes are involved in the activation of smaller alkanes (405–423 kJ mol
1, > Table 1) or even methane (439 kJ mol
1), the formation of the individual substrate radicals species from a glycyl-radical appears to be thermodynamically impossible. However, BSS and probably all other glycyl radical enzymes involved in anaerobic hydrocarbon degradation have a dimeric composition (Leuthner et al., 1998; Selmer et al., 2005), which opens the possibility that exergonic partial reactions of one monomer are coupled to endergonic reactions in the other monomer. Such a scenario has been proposed for the dual-stroke engine mechanism of methyl-coenzyme M reductase, which is involved in methane formation and anaerobic methane oxidation (Thauer and Shima, 2008) (see 2.3).

3.3

Reverse Methanogenesis

Only for a few years clear evidence is available that anaerobic methane oxidation is of important ecological relevance (Boetius et al., 2000; Niemann et al., 2006; Raghoebarsing et al., 2006). The methane-oxidizing organisms discovered first were archaea affiliated with the

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Anaerobic Degradation of Hydrocarbons

. Figure 3 Proposed radical species involved in catalytic turnover of benzylsuccinate synthase.

order Methanosarcinales that apparently form syntrophic associations with sulfate-reducing Deltaproteobacteria (Knittel et al., 2005; Nauhaus et al., 2005). However, the molecular basis of the required interspecies electron transfer of this syntrophic metabolism is still unknown. Biochemical analysis of anaerobic methane oxidizing microbial mats from the Black Sea revealed high concentrations (10% of total protein) of two Ni-containing methyl-coenzyme

Anaerobic Degradation of Hydrocarbons

15

M reductases (MCR), (Kru¨ger et al., 2003). This enzyme is usually involved in methanogenesis and catalyzes the formation of methane and a heterodisulfide from methyl-coenzyme M and coenzyme B. Surprisingly, the two types of MCR contained Ni-cofactors (F430) with different molecular masses. One of the cofactors was shown to contain an additional 172-methylthio-residue (+46 Da), which is missing in usual F430 cofactors (Thauer and Shima, 2008). Other differences between both MCRs were found; however, so far no rationale exists that assigns modifications in MCR enzymes to specific roles in methane formation/ oxidation. Only very recently evidence has been provided for the potential oxidation of methane coupled to denitrification (Raghoebarsing et al., 2006; Ettwig et al., 2008). The enriched culture consisted mainly of bacteria belonging to a novel phylum. So far, no experimental data about the enzymes involved have been obtained. However, thermodynamic considerations would not exclude a glycyl-radical enzyme-catalyzed initial addition of methane to fumarate yielding methylsuccinate (Thauer and Shima, 2008). In recent reviews by Thauer and Shima, the properties of methyl-coenzyme M reductases involved in methane formation/anaerobic methane oxidation and the thermodynamics/ kinetics of anaerobic methane oxidation were excellently summarized and are not presented here in detail (Shima and Thauer, 2005; Thauer and Shima, 2008). Briefly, reverse methanogenesis is thermodynamically realistic under in vivo conditions at rates as high as 0.1 U mg
1 protein. A possible explanation why different enzymatic mechanisms may be involved in anaerobic methane oxidation coupled to sulfate reduction (reversed methanogenesis) or nitrate reduction (possibly fumarate addition) might be the highly different energy yields as indicated by the following equations (Thauer and Shima, 2008). þ
CH4 þ SO2
4 þ H ! CO2 þ HS þ 2H2 O

DG0 ¼
21 KJ mol
1 þ 5 CH4 þ 8NO
3 þ 8H ! 5CO2 þ 4N2 þ 14H2 O

DG0 ¼
765 KJ per mol CH4 As a consequence, the proposed highly endergonic partial reactions in glycyl-radical enzymes such as the transient formations of the thiyl or benzyl radical are apparently energetically too costly when methane oxidation is coupled to sulfate reduction; in contrast, they appear to be negligible when coupled to denitrification.

3.4

Methylation and Carboxylation

There is an ongoing debate about the initial reactions involved in anaerobic degradation of benzene and naphthalene. In the case of naphthalene degradation, a first convincing evidence for the initial methylation to 2-methylnaphtahlene was obtained in a sulfate-reducing enrichment culture (Annweiler et al., 2002; Safinowski and Meckenstock, 2006). The methylated intermediate is then further converted by addition to fumarate in analogy to anaerobic toluene degradation, finally yielding 2-naphthoic acid (Safinowski and Meckenstock, 2004), (> Fig. 4a). The corresponding metabolites were identified and some of the proposed enzyme activities involved were determined in vitro. The enzyme involved in the proposed

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Anaerobic Degradation of Hydrocarbons

. Figure 4 Alternative activation reactions proposed for anaerobic degradation of naphthalene (a) and benzene (b).

methylation reaction is unknown; a likely candidate might be a B12-containing methyltransferase. As an alternative reaction, the direct carboxylation of naphthalene to 2-naphthoic acid has been discussed based on data obtained from labeling studies with [13/14C-] bicarbonate (Zhang and Young, 1997; Zhang et al., 2000), (> Fig. 4b). Additional evidence for an initial activation of naphthalene by carboxylation has recently been provided in studies with two different marine sulfate-reducing, naphthalene-degrading bacteria (Musat et al., 2008). When grown on naphthalene, these organisms metabolized 2-methylnaphthalene only after a lag phase and a glycyl radical enzyme-like protein putatively involved in methyl-group activation was only found to be expressed during growth on 2-methylnaphthalene but not during growth on naphthalene. Anaerobic degradation of benzene has been demonstrated in sulfate-reducing, Fe(III)reducing, and nitrate-reducing cultures, and even in a methanogenic consortium (Kunapuli

Anaerobic Degradation of Hydrocarbons

15

et al., 2008; Musat and Widdel, 2008; Ulrich et al., 2005 and references therein). Recently, two denitrifying Azoarcus strains were described as benzene-degrading pure cultures (Kasai et al., 2006, 2007), but the initial activation reaction of benzene still remains elusive. The dissociation energy of the C–H-bond of benzene is even higher than that of methane, which implies that a yet-unknown enzymatic reaction is involved. The mechanisms proposed were hydroxylation by water, methylation, or carboxylation of the ring. The occasionally reported hydroxylation of benzene is now rather assigned to an artificial side reaction of hydroxyl radicals, which are accidentally formed when traces of oxygen contaminate strictly anaerobic enrichment cultures in the presence of reducing agents (Kunapuli et al., 2008). In contrast, evidence for an initial methylation to toluene has been obtained (Ulrich et al., 2005) (> Fig. 4b). A recent study in a Fe(III)-respiring enrichment culture using 13C-benzene and 13 C-bicarbonate combined with a careful analysis of the time-dependent incorporation of the label in the product benzoic acid provided initial evidence for a direct carboxylation of benzene (Kunapuli et al., 2008). The concept of benzene carboxylation was supported by the fact that the Fe(III)-respiring enrichment culture degraded benzene but neither toluene nor phenol (Kunapuli et al., 2008; Musat and Widdel, 2008). The two latter compounds would represent intermediates in case of initial methylation (toluene) or hydroxylation reactions (phenol). The nature of the putative methylases/carboxylases involved in benzene/and or naphthalene degradation is unknown. The only similar carboxylation reaction known in nature is catalyzed by the key enzyme in anaerobic phenol degradation, phenylphosphate carboxylase, which converts its substrate to 4-hydroxybenzoate and inorganic phosphate (Schu¨hle and Fuchs, 2004). However, benzene and naphthalene cannot be directly activated by an ATP-dependent phosphorylation, which is necessary for phenol carboxylation. Thus, if carboxylation indeed represents the first step in benzene or naphthalene degradation, the putative carboxylases have to operate via a different mechanism. Anaerobic carboxylation of hydrocarbons has also been reported as initial step in the degradation of alkanes by some sulfate-reducing strains (Callaghan et al., 2006; So et al., 2003), but such reports need to be interpreted cautiously as incorporation of labeled carbon dioxide is not a true proof of the assumed carboxylation reactions.

4

Research Needs

The anaerobic metabolism of hydrocarbons is still a treasure chamber of novel, only poorly understood enzymatic reactions. These comprise the initial reactions involved in benzene and naphthalene degradation as well as in the anaerobic methane oxidation coupled either to sulfate reduction (via reverse methanogenisis) or to nitrate reduction (addition to fumarate?). Furthermore, almost nothing is known about the enzymology involved in the degradation of polycyclic aromatic or alicyclic hydrocarbons. Among all of the proposed glycyl-radical enzymes involved in the degradation of alkanes and aromatic hydrocarbons, only benzylsuccinate synthase has been studied in some detail. Nevertheless, so far no structure is available for any alkylsuccinate forming enzyme of the glycyl radical family. Studying the structure–function relationships of the enzymatic reactions involved in C–H-bond activation without oxygen not only will enable insights into novel biochemical processes but may also open a door for applications in biotechnology, bioremediation, and ecophysiology.

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Anaerobic Degradation of Hydrocarbons iron-sulfur/heme enzyme. J Biol Chem 276: 21381–21386. Knittel K, Losekann T, Boetius A, Kort R, Amann R (2005) Diversity and distribution of methanotrophic archaea at cold seeps. Appl Environ Microbiol 71: 467–479. Krieger CJ, Roseboom W, Albracht SP, Spormann AM (2001) A stable organic free radical in anaerobic benzylsuccinate synthase of Azoarcus sp. strain T. J Biol Chem 276: 12924–12927. Kru¨ger M, et al. (2003) A conspicuous nickel protein in microbial mats that oxidize methane anaerobically. Nature 426: 878–881. Kunapuli U, Griebler C, Beller HR, Meckenstock RU (2008) Identification of intermediates formed during anaerobic benzene degradation by an ironreducing enrichment culture. Environ Microbiol 10: 1703–1712. Leuthner B, et al. (1998) Biochemical and genetic characterization of benzylsuccinate synthase from Thauera aromatica: a new glycyl radical enzyme catalysing the first step in anaerobic toluene metabolism. Mol Microbiol 28: 615–628. Li L, Marsh EN (2006) Mechanism of benzylsuccinate synthase probed by substrate and isotope exchange. J Am Chem Soc 128: 16056–16057. Lieberman RL, Rosenzweig AC (2004) Biological methane oxidation: regulation, biochemistry, and active site structure of particulate methane monooxygenase. Crit Rev Biochem Mol Biol 39: 147–164. McIntire W, Hopper DJ, Singer TP (1985) p-Cresol methylhydroxylase. Assay and general properties. Biochem J 228: 325–335. McLeod MP, Eltis LD (eds.) (2008) Genomic insights into the aerobic pathways for degradation of organic pollutants. Norfolk: Academic Press. Morasch B, Schink B, Tebbe CC, Meckenstock RU (2004) Degradation of o-xylene and m-xylene by a novel sulfate-reducer belonging to the genus Desulfotomaculum. Arch Microbiol 181: 407–417. Mu¨ller JA, Galushko AS, Kappler A, Schink B (2001) Initiation of anaerobic degradation of p-cresol by formation of 4-hydroxybenzylsuccinate in Desulfobacterium cetonicum. J Bacteriol 183: 752–757. Musat F, et al. (2008) Anaerobic degradation of naphthalene and 2-methylnaphthalene by strains of marine sulfate-reducing bacteria. Environ Microbiol, 11(1): 209–219. Musat F, Widdel F (2008) Anaerobic degradation of benzene by a marine sulfate-reducing enrichment culture, and cell hybridization of the dominant phylotype. Environ Microbiol 10: 10–19. Nauhaus K, Treude T, Boetius A, Kruger M (2005) Environmental regulation of the anaerobic oxidation of methane: a comparison of ANME-I and

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ANME-II communities. Environ Microbiol 7: 98–106. Niemann H, et al. (2006) Novel microbial communities of the Haakon Mosby mud volcano and their role as a methane sink. Nature 443: 854–858. Peters F, Heintz D, Johannes J, van Dorsselaer A, Boll M (2007) Genes, enzymes, and regulation of para-cresol metabolism in Geobacter metallireducens. J Bacteriol 189: 4729–4738. Qiao C, Marsh EN (2005) Mechanism of benzylsuccinate synthase: stereochemistry of toluene addition to fumarate and maleate. J Am Chem Soc 127: 8608–8609. Rabus R, et al. (2001) Anaerobic initial reaction of n-alkanes in a denitrifying bacterium: evidence for (1-methylpentyl)succinate as initial product and for involvement of an organic radical in n-hexane metabolism. J Bacteriol 183: 1707–1715. Raghoebarsing AA, et al. (2006) A microbial consortium couples anaerobic methane oxidation to denitrification. Nature 440: 918–921. Safinowski M, Meckenstock RU (2004) Enzymatic reactions in anaerobic 2-methylnaphthalene degradation by the sulphate-reducing enrichment culture N 47. FEMS Microbiol Lett 240: 99–104. Safinowski M, Meckenstock RU (2006) Methylation is the initial reaction in anaerobic naphthalene degradation by a sulfate-reducing enrichment culture. Environ Microbiol 8: 347–352. Schu¨hle K, Fuchs G (2004) Phenylphosphate carboxylase: a new C-C lyase involved in anaerobic phenol metabolism in Thauera aromatica. J Bacteriol 186: 4556–4567. Selmer T, Pierik AJ, Heider J (2005) New glycyl radical enzymes catalysing key metabolic steps in anaerobic bacteria. Biol Chem 386: 981–988. Shima S, Thauer RK (2005) Methyl-coenzyme M reductase and the anaerobic oxidation of methane in methanotrophic Archaea. Curr Opin Microbiol 8: 643–648. So CM, Phelps CD, Young LY (2003) Anaerobic transformation of alkanes to fatty acids by a sulfate-reducing bacterium, strain Hxd3. Appl Environ Microbiol 69: 3892–3900. Szaleniec M, Hagel C, Menke M, Nowak P, Witko M, Heider J (2007) Kinetics and mechanism of oxygenindependent hydrocarbon hydroxylation by ethylbenzene dehydrogenase. Biochemistry 46: 7637–7646. Thauer RK, Shima S (2008) Methane as fuel for anaerobic microorganisms. Ann N Y Acad Sci 1125: 158–170. Ulrich AC, Beller HR, Edwards EA (2005) Metabolites detected during biodegradation of 13C6-benzene in nitrate-reducing and methanogenic enrichment cultures. Environ Sci Technol 39: 6681–6691. Verfu¨rth K, Pierik AJ, Leutwein C, Zorn S, Heider J (2004) Substrate specificities and electron paramagnetic resonance properties of benzylsuccinate

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synthases in anaerobic toluene and m-xylene metabolism. Arch Microbiol 181: 155–162. Widdel F, Rabus R (2001) Anaerobic biodegradation of saturated and aromatic hydrocarbons. Curr Opin Biotechnol 12: 259–276. 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.

Zhang X, Sullivan ER, Young LY (2000) Evidence for aromatic ring reduction in the biodegradation pathway of carboxylated naphthalene by a sulfate reducing consortium. Biodegradation 11: 117–124. Zhang X, 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.

16 The Role of Metals I. Bertini1,2,* . A. Rosato1,2 1 Magnetic Resonance Center (CERM), University of Florence, Sesto Fiorentino, Italy 2 Department of Chemistry, University of Florence, Sesto Fiorentino, Italy *[email protected] [email protected]

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1026 2 Manganese . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1027 3 Iron . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1027 4 Cobalt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1034 5 Nickel . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1036 6 Copper . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1037 7 Zinc . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1038 8 Molybdenum . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1039 9 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1040

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_72, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Metal ions are involved in the catalysis of many reactions that are crucial for the biodegradation of hydrocarbons. This is particularly true when the reaction involves redox chemistry, as transition metal ions have more than one oxidation state available under physiological conditions and can provide fast kinetics of electron transfer. They consequently constitute ideal components in the active site of metalloenzymes catalyzing redox reactions. The unpaired electrons available on metal ions are also important to provide a controlled manner to activate molecular dioxygen, when it acts as the oxidant on the hydrocarbon substrate. Some instances where metal ions participate in catalysis but are not redox active are also discussed. For each relevant transition metal ion, we describe the role in the best characterized enzymatic pathways and, whenever possible, try to present considerations on the reaction mechanisms that are of general interest.

1

Introduction

Hydrocarbons are often relatively inert compounds, due to the high energy of the C-C and C-H bonds as well as, when relevant, to the stability of aromatic systems. This has required that the microorganisms adapted their enzymatic portfolio to activate these bonds in a controlled manner. As many of the activation reactions involve redox chemistry, metalloenzymes have been an optimal (and therefore, widespread) choice in several instances. In addition, it is important to keep in mind that in many microbial:hydrocarbon interactions, a crucial role is played by molecular dioxygen, which is the strongest commonly available oxidant. However, the reaction of dioxygen with hydrocarbons normally features unfavorable kinetics and therefore is slow. This constitutes an additional determinant for the spread of metalloenzymes in microbes that metabolize hydrocarbons. Indeed, metal ions harboring unpaired electrons can efficiently activate molecular dioxygen and thus enable otherwise spin-forbidden reactions (Bertini et al., 2006). Overall, metal ions, as discussed in detail in this chapter, can play various roles to facilitate this ensemble of processes (Andreini et al., 2008), typically, but not exclusively, by activating the substrate (hydrocarbon), or the oxidant, or both, and therefore speeding up a slow but thermodynamically favorable reaction. Several metals play a role in the metabolism of hydrocarbons: manganese, iron, cobalt, nickel, copper, zinc, molybdenum can be involved in different stages of the process, also depending on the exact nature of the substrate (aromatic or aliphatic, long- or short-chain, etc) and on the specific system under consideration. Unsurprisingly, some solutions are more widespread than others, reflecting the compromise imposed by the chemistry of hydrocarbons on one hand and by the conditions needed to sustain microbial life on the other (e.g., aqueous environment, neutral pH, relatively low temperature). Bioinformatic tools can be successfully applied to reconstruct in silico metalloenzymatic pathways in organisms whose genome sequence is available (Bertini et al., 2007; Cavallaro et al., 2008). Finally, it is worth noting that a database specifically focusing on the mechanisms of the catalysis mediated by metalloenzymes has recently become available (Metal-MACiE, http:// www.ebi.ac.uk/thornton-srv/databases/Metal_MACiE/). The database describes, among other things, a significant variety of reactions whose substrate is a hydrocarbon. Only metalloenzymes of known three-dimensional structure are taken into account. For enzymes contained in the database, the relevant Id will be given. For enzymes not in Metal-MACiE,

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16

EC numbers are from the BRENDA enzyme database (http://www.brenda-enzymes.info/) (Barthelmes et al., 2007).

2

Manganese

Manganese plays a role in the biotransformation of hydrocarbons by manganese peroxidase (MnP, EC 1.11.1.13), an extracellular enzyme that is produced by several fungi, of which the peroxidase from Phanerochaete chrysosporium is probably one of the best characterized (Gold et al., 2000). In these systems, hydrogen peroxide performs a two-electron oxidation of the heme moiety to generate a iron(IV) = O/porphyrin cation species that in turn oxidizes manganese(II) to manganese(III). The latter is then chelated by a small organic molecule, such as oxalate, and diffuses away from the protein (Banci et al., 1998). The manganese(III) complex then encounters the substrate, which physiologically is lignin or a humic acid but for our purposes is an aromatic xenobiotic hydrocarbon (Eibes et al., 2007, 2006; Steffen et al., 2003), and performs a one-electron oxidation thereby generating a radical product. This product reacts spontaneously further. A reactive radical mediator can also be involved in the process, i.e., the manganese(III) complex does not oxidize directly the xenobiotic but instead acts on an intermediate organic compound, such as a thiol (Gold et al., 2000). In MnP, thus the manganese metal ion cycles between the +2 and +3 oxidation states and attacks a xenobiotic substrate directly or through an organic radical mediator (Wong, 2008). Manganese(II) is present as the physiologically relevant metal ion in a restricted number of extradiol ring-cleaving dioxygenases (Boldt et al., 1995; Que and Reynolds, 2000) (EC 1.13.11.2). These enzymes catalyze the cleavage of dihydroxybenzene rings with incorporation of both atoms from molecular dioxygen to yield muconate semialdehyde products. They more commonly contain a iron(II) ion in their active site. A thermostable, manganese (II)-dependent 2,3-dihydroxybiphenyl 1,2-dioxygenase, BphC_JF8, has also been reported (Hatta et al., 2003). The metal coordination is the same in two structurally characterized manganese(II) and iron(II)-dependent enzymes (MndD and homoprotocatechuate 2,3-dioxygenase, HPCD, respectively) (Vetting et al., 2004), which have a sequence identity as high as 83%. In spite of this striking sequence similarity, the two proteins are capable of selecting the native metal ion when the recombinant expression strain is grown in the presence of an equal concentration of manganese and iron in the medium. Through an elegant combination of metal substitution, kinetic studies and crystallographic characterization, it has been shown that the reaction mechanism is independent of the identity of the metal ion (Emerson et al., 2008). The role of manganese(II) in MndD is thus the same as that of iron(II) in HPCD, which has been characterized in detail (see next section). Computational studies suggest that even though the activation barrier is the same, the rate limiting step of the catalytic cycle may differ in the two enzymes MndD and HPCD (Georgiev et al., 2008).

3

Iron

Iron is probably the most important metal for the metabolism of hydrocarbons in microbial system. Iron is present in the active site of both mono- and dioxygenases (Bertini et al., 2006). Monooxygenases catalyze the addition of one atom from molecular dioxygen to the substrate, which thus gets hydroxylated. The second oxygen atom is reduced to water. Monooxygenases

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can contain iron in the form of non-heme iron or within a heme cofactor (Groves, 2006). Soluble methane monooxygenases (sMMO, EC 1.14.13.25) contains a di-iron center (Hakemian and Rosenzweig, 2007). The enzyme is in the fully oxidized form (iron(III)-iron (III)) in its resting state, and gets reduced to iron(II)-iron(II) at the expenses of one molecule of NADH (Lipscomb, 1994). The full reaction therefore is: RH þ O2 þNADH þ Hþ ! ROH þ H2 O þ NADþ The reduction of the diiron site is accompanied by a rearrangement of the coordination sphere of the two metal ions, both of which lower their coordination number from six to five. The reduced center has therefore the capability to interact with molecular dioxygen and reduce it to peroxide upon donation of one electron per iron ion. Upon uptake of two H+ ions, the peroxo-bridged species reacts further and one more electron per iron ion is transferred to the two oxygen atoms, yielding the release of a water molecule and a oxo-bridged iron(IV)-iron(IV) complex. This complex is not particularly stable and in the absence of substrate decays more rapidly than it is formed. When the substrate (not only methane but also other alkanes, alkenes, aromatics, and halogenated hydrocarbons (Colby et al., 1977)) is present, the oxo-bridged iron (IV)-iron(IV) complex can break homolytically a C-H bond and then rapidly hydroxylate the organic radical consequently generated. In this process the two iron ions are reduced from iron(IV) to iron(III). In sMMO, the two iron ions therefore act synergistically in activating molecular dioxygen by converting it to peroxide. Moreover, each of them provides an additional electron to the two oxygen atoms to reduce them to the 2 oxidation state. The resulting iron (IV)-iron(IV) species acts then as an acceptor of an electron pair, which is removed one electron at a time from the substrate. An analogous mechanism is operative in other monooygenases such as toluene monooxygenase (Murray et al., 2007; Murray and Lippard, 2007; > Fig. 1). Similar concepts apply to heme-containing monoxygenases, to which cytochromes P450 belong. Cytochromes P450 catalyze the hydroxylation (as well as of many other reactions) of small, hydrophobic substrates using a heme-iron as cofactor (Groves, 2006) (MACiE Id = M0133, EC 1.14.15.1), according to the reaction scheme of (1). In the resting state, the enzyme is oxidized (iron(III)), low-spin, with the iron ion exacoordinate by a water molecule and a protein cysteinate residue. Here, binding of the substrate occurs first, which triggers the release of the water molecules from the cavity, hence making the iron(III) ion pentacoordinate, high spin (Schenkman et al., 1967; Tsai et al., 1970). For several cytochrome P450 enzymes, this results in an increase of the iron ion reduction potential (Daff et al., 1997; Isin and Guengerich, 2008; Sharrock et al., 1973; Sligar and Gunsalus, 1976). A linear freeenergy relationship between the formal potential (E0 ) observed versus the free-energy change of the ferric spin state equilibrium has been found for two different cytochrome P450‘s, including one human (Das et al., 2007). After the aforementioned change in coordination, a partner ferredoxin transfers one electron yielding a iron(II) pentacoordinate site. In analogy to the case of sMMO, this site can now interact with molecular dioxygen and reduce it to superoxide first, upon oxidation of the iron ion, and then to peroxide, upon injection of a second electron by the partner. Similarly to sMMO again, addition of two H+ ions and a further two-electron reduction of the peroxide leads to heterolytic cleavage of the O-O bond, with release of a water molecule. One of the latter two electrons is donated by the iron(III) ion and the other by the porphyrin moiety, leading to formation of a so-called ferryl (iron (IV) = O) species and of a porphyrin radical. This species can break homolytically a C-H bond of the substrate, and the nascent radical ‘‘rebounds’’ with the Fe-OH moiety to yield the hydroxylated product leaving the enzyme in the iron(III) state > Fig. 2. The mechanisms of

The Role of Metals

. Figure 1 Reaction scheme of iron-dependent MMO.

. Figure 2 Reaction scheme of cytochrome P450.

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action of the diiron cluster contained in a variety of monooxygenases and the iron-porphyrin moiety of cytochromes P450 bear some interesting similarities. Indeed, these systems can be viewed as containing a iron(II) ion plus an electron donor. The pentacoordinate iron(II) moiety interacts with molecular dioxygen yielding formation of an iron(III)-superoxide complex that accepts a further electron (either from the other metal ion or from an external source) to form the key peroxo intermediate. At this point, the iron(III) ion and the electron donor (the second iron(III) ion in the diiron cluster or porphyrin, respectively) promote heterolytic cleavage of the O-O bond, releasing water and forming a iron(IV) species plus the oxidized electron donor. The iron(IV) species can homolytically cleave a C-H bond, restoring the reduced electron donor and forming a Fe(IV)-hydroxo intermediate. A subsequent rebound step yields the hydroxylated product and the oxidized enzyme. The high reduction potential of the iron(IV) species is crucial to enable the latter reaction, and is modulated by the coordination and environment of the iron ion (Que, 2007). A significant difference is instead in the fact that diiron monooxygenases bind molecular dioxygen before the substrate is present, whereas the iron(II) site in cytochromes P450 becomes available only after the substrate enters the enzyme cavity thereby allowing the iron ion to be reduced > Fig. 2. A large variety of enzymes relevant for the present book contain mononuclear non-heme iron sites. These enzymes most commonly catalyze the insertion of both atoms of molecular dioxygen into the substrate (Koehntop et al., 2005). The cis-dihydroxylation of arenes catalyzed by Rieske dioxygenases constitutes the first step in the biodegradation of aromatic molecules by soil bacteria and leads in two subsequent steps to the formation of catechols. In naphthalene dioxygenase (MACiE Id = M0130, EC 1.14.12.12), the alpha subunit of the enzyme contains a mononuclear iron site and a Rieske cluster (Karlsson et al., 2003; Kauppi et al., 1998). The mononuclear site is reduced in the resting state and pentacoordinate, while the cluster is oxidized. The enzyme cycle starts with a one-electron reduction of the cluster. Then, binding of the substrate in the enzyme cavity allows molecular dioxygen to diffuse to the iron(II) ion and react with it. An electron is simultaneously delivered from the Rieske cluster, yielding a iron(III)-peroxo adduct. This species reacts directly (or via a transient iron(V) species according to some models (Chen et al., 2002)) with the substrate yielding, upon the subsequent uptake of one electron, the di-hydroxylated species and the enzyme in its resting state. The reaction catalyzed is as shown in the > Figs. 3 and 4. Dihydroxybenzene rings are the substrate of ring-cleaving dioxygenases. Extra-diol dioxygenases acting on ortho-catechols yield muconate semialdehyde products, whereas intra-diol dioxygenases yield cis, cis muconates (Vaillancourt et al., 2006). Other substrates include, for example, para- catechols such as gentisate (Werwath et al., 1998) and typically yield products similar to those of extra-diol dioxygenases > Fig. 5. In the resting state of catechol 2,3-dioxygenase (MACiE Id = M0034, EC 1.13.11.2), the mononuclear iron site is reduced and penta- (Sato et al., 2002; Shu et al., 1995) or

. Figure 3 Reaction catalyzed by Rieske dioxygenases.

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. Figure 4 Reaction scheme of Rieske dioxygenases.

. Figure 5 Comparison of the products of extra- and intra-diol dioxygenases.

exa-coordinate (Kovaleva and Lipscomb, 2007). The substrate binds to the metal as a bidentate, doubly negatively charged ligand, removing two or three water molecules/hydroxide ions leaving in all cases a pentacoordinate iron(II) ion. At this point, molecular dioxygen can be bound by the metal. Notably, binding is not associated with a redox reaction. Instead, the dioxygen molecule acts as a nucleophile on the C2 atom (i.e., one of the two hydroxylated carbons), leading to the formation of an alkylperoxo species. A series of molecular rearrangements eventually leads to the cleavage of the O-O bond and the formation of the product (2-hydroxymuconate semialdehyde), which dissociates from the metal ion. The x-ray structures of three intermediates in the cycle catalyzed by homoprotocatechuate 2,3-dioxygenase (HPCD) suggested that the substrate initially donates some electron density to the dioxygen

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molecule to form a superoxide anion, which binds side-on (i.e., with both oxygen atoms within bond distance) to the iron(II) ion, and an alkyl cation radical (Kovaleva and Lipscomb, 2007). The two then react to form the aforementioned alkylperoxo species, which has been structurally characterized (Kovaleva and Lipscomb, 2007). As mentioned in the preceding section, it has been demonstrated that HPCD has the same mechanism as the manganesedependent enzyme MndD (Emerson et al., 2008). Because of the significant different in reduction potential between the iron(III)/iron(II) and the manganese(III)/manganese(II) pair in the otherwise identical metal binding site, and given the results from the crystallography study of HPCD, it appears that the role of the metal in extra-diol ring-cleaving dioxygenases is only that of suitably orienting the two substrates to permit their interaction and, possibly, of electrostatically stabilizing the intermediates along the reaction. Intra-diol ring-cleaving dioxygenases (e.g., protocatechuate 3,4-dioxygenase, EC 1.13.11.3) feature a mononuclear iron(III) site in the resting state. The higher oxidation state than in extra-diol dioxygenases is stabilized by the presence of two negatively charged tyrosinate ligands from the protein, in addition to two histidines and a water molecule, which compares to a coordination environment featuring two water molecules, two histidines and a monodentate carboxylate in extra-diol dioxygenases. The higher negative charge and stronger electron-donating character of tyrosinate in the former site stabilizes iron(III) versus iron(II). Similarly to what described in the preceding paragraph, the first event during the catalytic cycle is the binding of the substrate to the iron(III) ion as a bidentate ligand. The substrate replaces one water molecule and one tyrosinate. The iron complex thus formed has a partial iron(II)-substrate radical character that renders the substrate prone to attack by molecular dioxygen. In fact, the latter reacts with the complex, yielding an alkylperoxo intermediate occupying three of the six coordination sites of the iron(III) ion, analogously to the case of extra-diol dioxygenases. This species rearranges to cleave the O-O bond and generate the cis, cis muconate product. Here the role of iron ion is mainly to activate the substrate. Theoretical studies indicate that the metal is also key in overcoming the spin-forbidden nature of the molecular dioxygen reaction by acting on the spin state of an electron from the substrate to allow the second electron reduction of O2, while leaving an intermediate spin state (S = 3/2) on the iron(III) center (Pau et al., 2007; > Fig. 6). Rieske dioxygenases and extra-diol ring-cleaving dioxygenases share a common structural feature at the metal site: the iron(II) center is invariably coordinated by three protein residues, two His and one Asp or Glu, constituting one face of an octahedron, a recurring motif referred to as the 2-His-1-carboxylate facial triad (Koehntop et al., 2005; Que, 2000). The 2-His-1carboxylate facial triad serves as a monoanionic three-pronged platform for binding divalent metal ions. The three remaining sites on the opposite face of the octahedron are consequently available for exogenous ligands. In the resting state of the enzymes, two or all three of these sites are usually occupied by solvent molecules. During catalysis, they can accommodate both the substrate and molecular dioxygen. Notably, the iron(II) ion is relatively unreactive toward dioxygen before the substrate enters the enzyme active site. In other words, the metal center becomes poised to bind molecular dioxygen only when the substrate (and, where relevant, the cofactor(s)) is present in the active site, thereby promoting strong coupling between the reduction of dioxygen and the oxidation of substrate. A similar feature is observed also in the case of cytochromes P450 and, to some extent, of intra-diol ring-cleaving dioxygenases, suggesting that this can constitute an evolutionarily favorable strategy that avoids unnecessary, and potentially dangerous, binding and activation of dioxygen when the substrate is not readily available. As detailed above, the role of the iron ion in the various kind of mononuclear

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. Figure 6 Reaction schemes of extra- (bottom) and intra-diol (top) dioxygenases.

non-heme sites that are relevant for hydrocarbon degradation is quite differentiated, even within enzymes with the 2-His-1-carboxylate facial triad. This role can entail a variation of the oxidation state, or not. Regarding molecular dioxygen, a general key intermediate is the peroxo species either in the form of a peroxide complex with the metal or of an alkylperoxo intermediate, also coordinating the metal ion. In the Rieske dioxygenases, the peroxide complex is side-on (Karlsson et al., 2003) (as well as the superoxide ion in HPCD (Kovaleva and Lipscomb, 2007)), whereas it is end-on (i.e., only oxygen atom binds to the metal, the other being protonated) in cytochromes P450 (Schlichting et al., 2000). The selection between these two mechanisms presumably depends on the availability of a H+ donor and of suitable groups for H-bond interaction(s) within the enzyme active site. In the side-on mode, both oxygen atoms are equally activated thus favoring homolytic cleavage and dioxygenation (Kovacs, 2003). In the end-on mode, heterolytic cleavage and monoxygenation are favored. Up to this point, only iron enzymes using molecular dioxygen were mentioned. Fungi can produce extracellular haloperoxidases, some of which can catalyze also the hydroxylation of aromatic substrates using H2O2 as the oxidant (Ullrich and Hofrichter, 2007, 2005). These enzymes contain iron(III)-porphyrin in their resting state and bind hydrogen peroxide to give an intermediate analogous to that of cytochromes P450 after the second electron is received (i.e., the iron-(III)-peroxo state). From this point on, they follow the same reaction mechanism of cytochromes P450 (Ullrich and Hofrichter, 2007). In the anaerobic metabolism of hydrocarbons, a key enzyme containing iron in its active site is benzoyl-CoA reductase (BCR, EC 1.3.99.15) which catalyzes the reductive dearomatization of benzene rings, using the energy derived from the hydrolysis of ATP. BCR is composed by four subunits and contains three [4Fe-4S]2+ iron-sulfur clusters, of which one bridges subunits I and IV (Boll et al., 2000, 2001a). In the one-electron reduced protein, it is not possible to observe a single [4Fe-4S]+ cluster, but rather a mixture of interacting species (Boll et al., 2000, 2001a). Addition of MgATP to the reduced protein initiates its ATPase activity and triggers formation of a species having spin S = 7/2 (rather than the usual S = 1/2 state) that could be detected spectroscopically. It has been calculated that this transition lowers the reduction potential of the polymetallic

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. Figure 7 Steps in the reduction of benzoyl-CoA.

site by 0.5 V and brings it in the range suitable for reducing the substrate. Because the reduction of the benzene ring requires two electrons, which are transferred one at a time, the transition from S = 1/2 to S = 7/2 followed by the re-oxidation of the cluster by the substrate must be repeated twice per enzymatic cycle. Indeed, BCR binds two ATP molecules prior to the reduction of benzoyl-CoA (Mobitz et al., 2004). The reaction therefore proceeds via the following steps as shown in > Fig. 7. At each reaction steps the iron-sulfur cluster cycles between the + 2 and + 1 oxidation states. The difficult step is the transfer of the first electron to the aromatic substrate, which may be assisted by the simultaneous protonation at the para-position thereby avoiding the formation of a true radical anion intermediate. The first electron transfer could be further facilitated by partial protonation of the carbonyl oxygen. These two latter substrate-protein interactions would effectively raise the reduction potential of the aromatic substrate, therefore reducing the energy gap between the reductant (the [4Fe-4S]+ cluster in the S = 7/2 form) and the oxidant (the substrate). It is to be noted that the mechanism described above is relatively speculative, also given the absence of structural data for the enzyme.

4

Cobalt

Cobalt in enzymes is coordinated by a corrin ligand. One of the most studies corrinoids is cyanocobalamin, a member of the vitamin B12 family. Cobalt is involved in enzymes catalyzing respiratory dehalogenation of haloorganics, a process in which the latter compounds are used as terminal acceptors of electrons in the respiratory process. A possible general mechanisms for the dehalorespiring reductive dehalogenases involves the known (Banerjee and Ragsdale, 2003) ability of cobalt in cobalamins to form Co-C bonds. Reduced (cobalt(I)-containing) cobalamin would transfer one electron to a compound such as tetrachloroethylene, promoting release of a chloride ion. The nascent organic radical would combine with the cobalamin cofactor, receiving a further electron from the metal ion. Note that there is no information on the coordination of the cobalt ion throughout the catalytic cycle, except that it is tetra-coordinate in the cobalt(II) state (Schumacher et al., 1997). Indeed, the typical binding motif for corrinoids found in vitamin B12-dependent mutases and methionine synthases is not contained in tetrachloroethene reductive dehalogenase (Maillard et al., 2003). Protonation leads to the release of trichloroethylene, leaving a cobalt(III) ion that is subsequently re-reduced to the resting enzyme (Holliger et al., 1999). Another possible mechanism has been proposed (Schumacher et al., 1997), e.g., with chlorinated aromatics, where the reduced cobalamin would transfer its electron to the substrate that then forms an anion radical, which is rapidly protonated

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16

by the enzyme. The neutral radical would receive a further electron, which may or may not be provided by the cobalt(II) ion, to eliminate a chloride ion. In this second mechanism, the capability of cobalt to form Co-C bonds would not be exploited and cobalamin would function only as a reductant. The reaction catalyzed for the substrate tetrachloroethylene is: Cl2 C ¼ CCl2 þ2e þHþ ! Cl2 C ¼ CHCl þ Cl The mechanism involving formation of a Co-C bond has been challenged by the difficulty of producing a model Co(III)-trichlorovinyl complex (Kliegman and McNeill, 2008). Formation of an organometallic intermediate on the other hand is substantiated by the regioselectivity observed in the case of the dehalogenation of trichloroethylene (Glod et al., 1997), either in the form of a dichlorovinyl- or of a trichloroethyl-cobalt complex > Fig. 8. Cobalt also has a role in the activation of methanol for subsequent reduction to methane or oxidation to carbon dioxide. Methanol metabolism is initiated in methanogenic archaea by its reaction with coenzyme M (HS-CoM) to form methyl-coenzyme M (CH3-S-CoM) (Thauer, 1998). Methyl-coenzyme M is the central intermediate for the subsequent reactions. The enzyme catalyzing the formation of methyl-coenzyme M is methanol:coenzyme M methyltransferase (Mta, EC 2.1.1.90). The heterolytic cleavage of the inert methanol molecule is accomplished by a combined effect of the reduced cobalt(I) ion in the cofactor 5-hydroxybenzimidazolyl cobamide and of a nearby zinc(II) ion acting as an electrophile, as demonstrated by the three-dimensional structure of the enzyme (Hagemeier et al., 2006). The C-O bond in methanol is activated by the interaction of its hydroxyl group with the zinc

. Figure 8 Reaction scheme of reductive dehalogenase.

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(II) ion and can be attacked by cobalt(I), acting as a nucleophile. The heterolytic cleavage of the bond results in the formation of a methyl-cobalt(III) complex, to which some electron density is contributed by the formation of a coordination bond between the cobalt(III) ion and a nearby histidinate (His136) that happens simultaneously with methylation. This mechanism is consistent with the stereochemistry of the methyl group transfer from methanol to coenzyme M, which proceeds with retention (two times inversion) of stereo-configuration (Zydowsky et al., 1987). The axial ligand His136 is crucial for the catalytic reaction: indeed, the methylation rate of free cob(I)inamide (which lacks the imidazole base) with methanol catalyzed by MtaB is strongly enhanced by the presence of imidazole (Sauer and Thauer, 1999). The subsequent transfer of the methyl group from the organometallic complex to coenzyme M requires a rearrangement of the quaternary structure of Mta, which is a multisubunit enzyme (Hagemeier et al., 2006). The mechanism for this reaction is not known in detail > Fig. 9. The above data concur in describing the role of cobalt in the interaction of prokaryotes with hydrocarbons as a relatively strong reductant (the cobalt(II)/cobalt(I) couple in cyanocobalamin has E0 = 0.59 V (Schrauzer et al., 1968)), sometimes coupled to formation of organometallic complexes after heterolytic cleavage of one bond of the substrate.

5

Nickel

A key role of nickel is in reverse methanogenesis, i.e., the oxidation of methane carried out by prokaryota under anaerobic conditions (typically using sulphate as the oxidant (Boetius et al., 2000; Shima and Thauer, 2005); a nitrate reducing consortium has been identified in 2006 (Raghoebarsing et al., 2006)). The first step in the process would be carried out by a methyl-CoM reductase-like protein (MCR-like) catalyzing the reverse reaction with respect to the ‘‘ordinary’’ enzyme found in methanogens (Hallam et al., 2004). The prosthetic group putatively responsible for methyl transfer to the CoM cofactor is spectroscopically similar to F430 (Kruger et al., 2003) and contains a single nickel(II) ion. In the next step an enzyme similar to methyl-H4MPT:CoM methyl-transferase (N5-methyltetrahydromethanopterincoenzyme M methyltransferase), again working in the reverse direction, would be involved. The cofactor of the latter enzyme has not been identified; the enzyme contained in methanogens has a 50 -idroxybenzimidazolyl-cobamide cofactor, which contains cobalt. In the absence of any characterization of the MCR-like enzyme, it can only be hypothesized that it functions

. Figure 9 Reaction scheme of methanol:coenzyme M methyltransferase.

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by reverting the mechanism of MCR or through the initial formation of a Ni(III) hydride activating methane with the simultaneous release of H2 (Harmer et al., 2008).

6

Copper

Particulate methane monooxygenase (pMMO, EC 1.14.13.25) is an integral membrane copper-containing enzyme that converts methane to methanol (see scheme 1). The publication in 2005 of its three-dimensional structure has been a breakthrough for the understanding of its mechanisms of action (Lieberman and Rosenzweig, 2005), due to the difficulty of characterizing its metal content, organization, and location of the active site. pMMO is produced by the majority of methanotrophs and is more commonly found than the irondependent sMMO (Dumont and Murrell, 2005). In organisms that possess both forms, including Methylococcus capsulatus (Bath) and Methylosinus trichosporium OB3b, differential expression is regulated by the bioavailable copper concentration (Murrell et al., 2000). pMMO is more selective than sMMO toward alkanes and alkenes that are five carbons or less (Burrows et al., 1984). Three metal centers were identified in the crystal structure (Lieberman and Rosenzweig, 2005): a mononuclear copper center located in the soluble domain of the B subunit, 25 A˚ from the surface of the membrane; a dinuclear copper center in the B subunit located 21 A˚ from the mononuclear copper site; a zinc ion within the membrane, 19 A˚ from the dicopper center and 32 A˚ from the mononuclear copper site. The latter site probably houses a different metal ion in vivo (Hakemian and Rosenzweig, 2007); a recent X-ray characterization of Methylosinus trichosporium OB3b pMMO showed the presence of only two metal sites: a dinuclear copper center and a mononuclear copper center corresponding to the aforementioned zinc site (Hakemian et al., 2008). Other authors have suggested that a cluster of hydrophilic residues in the structure can bind a tri-copper cluster that has been removed from the crystallized protein by the purification procedure (Chan and Yu, 2008). The presence of the tri-copper cluster in the active enzyme has spectroscopic support (Chan et al., 2007). The uncertainty on the location of the active site and the chemical nature of the copperbased catalytic center clearly limits the possibility to reliably describe the mechanism of the enzyme and therefore to define the role of the copper ions. Because of the absence of an experimentally validated model for the structure of the tri-copper cluster, we will mention here only the mechanism proposed for the di-copper cluster seen in the crystal structure. The enzyme would bind molecular dioxygen in the copper(I)copper(I) state to yield a copper(II) copper(II)-peroxo species, in equilibrium with a copper(III)copper(III)-bis-oxo species (Balasubramanian and Rosenzweig, 2007). Injection of an electron might then generate the mixed valent bis(m-oxo)copper(II)copper(III) complex, which has been suggested to be more reactive (Yoshizawa and Shiota, 2006). This species would be responsible for the cleavage of the C-H bond of methane, formation of a non-radical intermediate, and recombination of OH and CH3. An alternative pathway involves oxidation of the mononuclear copper site to give a copper(III)–oxo complex, which would then act as the mixed-valence species described above. However, the formation of the copper(III)–oxo complex is predicted to be significantly less favorable from the thermodynamic point of view than the formation of the mixed valent bis (m-oxo)copper(II)copper(III) complex (Yoshizawa and Shiota, 2006). The copper-containing active site of pMMO functions similarly to the di-iron site of sMMO, by reducing molecular dioxygen to its final 2 oxidation state while generating metal

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The Role of Metals

species with a high oxidation state, which are then responsible for the cleavage of the C-H bond. However, it should be noted that despite the two mechanisms may appear similar, the cleavage of the C-H bond and formation of the product involves a concerted non-radical mechanism, as shown by the retention of the configuration of chiral substrates (Yu et al., 2003). Instead, sMMO involves a hydrogen-abstraction radical-rebound mechanism (see the section on Iron). Among copper enzymes, laccases (p-diphenol:dioxygen oxidoreductase, EC 1.10.3.2) have received attention for their potential in biotechnological applications and bioremediation, particularly for the removal of polycyclic aromatic hydrocarbons Rodriguez and Toca Herrera (2006). Laccases are extracellular multi-copper oxidases expressed mainly in plants and fungi that do not insert oxygen atoms in the substrate. They contain a tri-copper and a mononuclear copper center, which is close in space to the copper cluster (Piontek et al., 2002). Collectively, the four copper(I) ions of the resting enzyme can donate four electrons to molecular dioxygen and generate two molecules of water; the reduction takes place at the tri-copper site (Solomon et al., 1996). Each of the four oxidized copper(II) ions can then receive one electron from one molecule of substrate, either directly or through a chemical mediator, generating phenoxyl radical species that eventually get hydroxylated. The interaction of laccase with compounds such as 2,4-diamino-6-nitrotoluene, a possible metabolite of reduced 2,4,6-trinitrotoluene, leads to the formation of hydroquinones or benzoquinones (Dawel et al., 1997).

7

Zinc

Alcohol dehydrogenases (ADHs, EC 1.1.1.1) constitute a large family of enzymes responsible for the reversible oxidation of alcohols to aldehydes with the concomitant reduction of NAD+ or NADP+. These enzymes have been identified in eukaryotes and prokaryotes. Among other things, ADHs also support the growth of methylotrophs, oxidize alcohols and catalyse lignin degradation (Reid and Fewson, 1994). As an example, the xylB gene of the TOL plasmid pWW0 of Pseudomonas putida encodes a benzyl alcohol dehydrogenase, which is relevant in the pathway of transformation of toluene (and xylenes) into benzoate for subsequent degradation (Harayama et al., 1989). A closely related enzyme was found also in Acinetobacter calcoaceticus (Gillooly et al., 1998), which is able to use a large variety of aromatic compounds as the sole carbon and energy sources. The zinc(II) ion is not redox active; its role in alcohol dehydrogenases is that of coordinating the oxygen of the alcohol thereby increasing the acidity of the substrate and facilitating its deprotonation and the subsequent transfer of the hydride ion to NAD+ (Ramaswamy et al., 1999). The reaction catalyzed is as shown in > Fig. 10.

. Figure 10 Reaction catalyzed by benzyl-alcohol dehydrogenase.

The Role of Metals

8

16

Molybdenum

Besides one other type of molybdenum-containing cofactor, molybdenum is bound to a unique tricyclic pterin compound (molibdopterin) to form the Moco cofactor (Hille, 1996). The other type of cofactor is found only in bacterial nitrogenase, forming the so-called FeMocofactor that consists of two partial cubanes (MoFe3S3 and Fe4S3) which are joined by three bridging sulfurs (Howard and Rees, 2006). The members of the aldehyde oxidase (AO, EC = 1.2.3.1, METAL-MACiE Id M0105) family are cytoplasmic enzymes that catalyze the oxidation of a variety of aromatic and nonaromatic heterocycles and aldehydes, thereby converting them to the respective carboxylic acid. AO enzymes exclusively use molecular oxygen as electron acceptor. In AO’s the molybdenum(VI)-hydroxyl group performs a nucleophylic attack on the substrate with the help of a nearby base (Doonan et al., 2005). The concerted hydride transfer to the sulfur atom of the molybdenum(VI) = S group leads to formation of molybdenum(IV)-SH group, with the metal ion coordinated to the newly formed carboxylate group of the oxidized substrate. After dissociation of the product, two Fe2S2 clusters coordinated by the same protein chain relay two electrons to an acceptor, to restore the oxidized molybdenum(VI) cofactor. A related enzyme is 4-hydroxybenzoyl-CoA reductase (HBCR, EC = 1.3.99.20), which is involved in the anaerobic metabolism of aromatic compounds. HBCR catalyzes the reductive elimination of the phenolic hydroxyl group from the substrate (Brackmann and Fuchs, 1993; > Figs. 11 and 12). The reaction product, benzoyl-CoA, is the substrate of an iron-containing enzyme already discussed. With respect to AO, HBCR catalyses the reverse reaction, in which the substrate is reduced and a ferredoxin protein partner oxidized. Crucial to the functioning of the enzyme is the control of the reduction potential of the molybdenum ion in the Moco cofactor: the molybdenum(V)/molybdenum(IV) couple in HBCR has a potential as low as 500 mV (Boll et al., 2001b). The mechanism is still unclear, but it appears not to be a simple reversal of the general mechanism of members of the xanthine oxidase family (to which also AO belongs). Rather, it involves reactive radical species. The substrate coordinates to the molybdenum(IV) of the enzyme with its phenolic hydroxyl group. A one-electron reaction generates a radical anion and a molybdenum(V) species. The protonation of the para-carbon of the aromatic ring of the anion, possibly by the molybdenum(V)-SH moiety, promotes the homolytic cleavage of the C-O bond leaving a molybdenum(VI)-OH species, which is then reduced by two external electrons back to molybdenum(IV) (Boll et al., 2001b).

. Figure 11 Reaction catalyzed by HBCR.

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. Figure 12 Reaction scheme of HBCR.

9

Research Needs

As it is apparent also from the above data, there are two clear directions for future work on the role of metals in microbial:hydrocarbon interactions. One is that of obtaining structural characterization for the large variety of metalloenzymes for which some kinetic or functional data, or both, are available, but which lack details at the atomic level on the structure of the metal site and the role, or even identity, of key amino acids within the active site. Another outstanding need is that of extending the ensemble of known metalloenzymes with a biochemical characterization at any level, as there are clear indications that there are many more metalloenzymes in the environment than we presently can describe to a satisfactory extent in handbooks.

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17 Biochemistry and Molecular Biology of Methane Monooxygenase J. C. Murrell1 . T. J. Smith2 Department of Biological Sciences, University of Warwick, Coventry, UK [email protected] 2 Biomedical Research Centre, Sheffield Hallam University, Sheffield, UK

1

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1046 2 Biochemistry of Particulate Methane Monooxygenase (pMMO) . . . . . . . . . . . . . . . . . . . 1047 3 Biochemistry of Soluble Methane Monooxygenase (sMMO) . . . . . . . . . . . . . . . . . . . . . . . 1048 4 Molecular Biology and Regulation of Methane Monooxygenases . . . . . . . . . . . . . . . . . . 1049 5 Methanotrophs in Biocatalysis and Bioremediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1050 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1053

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_73, # Springer-Verlag Berlin Heidelberg, 2010

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Biochemistry and Molecular Biology of Methane Monooxygenase

Abstract: Methane oxidizing bacteria (methanotrophs) are a unique group of aerobic bacteria that can gain all of their carbon and energy requirements from methane. The enzymes that catalyze the first step in the bacterial methane oxidation pathway, the oxidation of methane to methanol, are called methane monooxygenases. These are remarkable enzymes because methane is chemically very stable and to convert methane to methanol chemically requires expensive catalysts, high temperatures and pressures. There are two types of methane monooxygenase that occur in methanotrophs, a membrane-bound, particulate methane monooxygenase and a cytoplasmic, soluble methane monooxygenase which belongs to a class of enzymes known as soluble di-iron monooxygenases. The expression of these enzymes in methanotrophs is often regulated by the availability of copper. The soluble methane monooxygenase has attracted significant attention and has considerable potential in biocatalysis and bioremediation since it can co-oxidize a very wide range of aliphatic and aromatic compounds, even though methanotrophs themselves do not grow on these compounds. We review here the biochemistry and molecular biology of both the particulate and soluble methane monooxygenases and their biotechnological potential.

1

Introduction

Methane-oxidizing bacteria (methanotrophs) are remarkable in being able to use the inert methane molecule to provide all of the chemical energy for the cell and also to synthesize the carbon building blocks for all of the macromolecules in the cell. They carry out the oxidation of methane via the enzyme methane monooxygenase (MMO) and subsequently use the same enzymes found in other aerobic Gram negative methylotrophic bacteria for further oxidation of methanol to formaldehyde, formate and carbon dioxide and for assimilation of carbon, at the oxidation level of formaldehyde, into cellular constituents (Anthony, 1982; Dalton, 2005; Hanson and Hanson, 1996; Trotsenko and Murrell, 2008). In methanotrophs there are two structurally and biochemically distinct forms of MMO, particulate (pMMO) and soluble methane monooxygenase (sMMO), which oxidize methane to methanol. pMMO is a copper-containing enzyme that is associated with unusual intracellular membranes found in type I methanotrophs as vesicular disks arranged in bundles throughout the cell and as paired peripheral layers in type II methanotrophs. sMMO is a cytoplasmic non-heme iron enzyme complex. The best characterized methanotrophs, Methylococcus capsulatus (Bath) (type I) and Methylosinus trichosporium OB3b (type II), can produce either form of MMO (reviewed in Murrell et al., 2000). The single factor known to govern expression of the two types of MMO in these organisms is the concentration of available copper (see later). At high copper-to-biomass ratio, pMMO is produced whereas the soluble form of the enzyme is expressed only when the copper-to-biomass ratio during growth is low (Stanley et al., 1983). Many methanotrophs such as the Type I methanotrophs Methylomonas methanica and Methylomicrobium album BG8 possess only pMMO and until recently the dogma was that all methanotrophs contained pMMO. Recently, however, the facultative type II methanotroph Methylocella silvestris has been shown to posses only the sMMO system and does not possess pMMO (Dedysh et al., 2005; Thiesen et al., 2005). These two families of MMOs share no detectable similarity in amino acid sequence or three-dimensional structure and are not evolutionarily related. It may be because methane is such a small and unfunctionalized substrate that both sMMO and pMMO are able to co-oxidize a range of

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hydrocarbons and chlorinated pollutants in addition to their natural substrate. Hence sMMO and pMMO have biotechnological potential that extends far beyond their ability to oxidize methane to methanol (see later).

2

Biochemistry of Particulate Methane Monooxygenase (pMMO)

pMMO is a copper-containing, membrane-associated enzyme (Nguyen et al., 1998; Smith and Dalton, 1989; Zahn and DiSpirito, 1996) and molecular ecology studies indicate that pMMO is probably responsible for most of the oxidation of methane carried out by aerobic methanotrophs in the environment (reviewed in McDonald et al., 2008). Being a membrane protein, the biochemistry of pMMO has lagged behind that of sMMO largely due to problems in solubilizing the pMMO away from membranes and purifying it in active form. The use of dodecyl-b-D-maltoside as detergent (Smith and Dalton, 1989), however, allows recovery of activity after solubilization and subsequent development of purification protocols have allowed the enzyme to be purified in an active form. Active preparations of pMMO generally contain three polypeptides, of about 49, 27 and 22 kDa. The 27-kDa subunit can be labeled by the inhibitor acetylene (a suicide substrate for both pMMO and sMMO) and until recently it was thought that the active site resided on this subunit. More recent structural studies suggest, however, that the active site may reside on the 45 kDa subunit or indeed may be shared between more than one subunit (reviewed in Hakemian and Rosenzweig, 2007). The 49, 27 and 22-kDa components are encoded by the genes pmoA, B and C, respectively, which are multicopy genes (see below) that are induced in response to growth of methanotrophs at high copper-to-biomass ratio. The recent determination of a crystal structure of pMMO, albeit with protein of rather low activity, shows that the enzyme has an (abg)3 stoichiometry and gives the first indication of the atomic-resolution structure of the enzyme (Lieberman and Rosenzweig, 2005). Single-particle analysis and associated biochemical studies have indicated that native pMMO forms a complex with methanol dehydrogenase, which may supply electrons to the enzyme (Kitmitto et al., 2005; Myronova et al., 2006). Whilst all active preparations of pMMO contain copper, the numbers and roles of copper ions in the active form of the enzyme continue to be hotly debated, as does the possible involvement of iron in the metal centres of the enzyme and the possibility that the iron may (as in sMMO) be organized into diiron centres (Martinho et al., 2007 reviewed in Hakemian and Rosenzweig, 2007). Little is currently known about the catalytic cycle of pMMO. Retention of stereochemistry is observed during oxygenation of certain chiral alkanes and so the mechanism of C-H bond breakage is likely to be concerted (rather than involving radical or cation intermediates). It will be interesting to see what similarities there are between the catalytic mechanism of pMMO and sMMO, which catalyse the same reaction within such different enzyme environments. The substrate profile of pMMO is very much narrower than that of sMMO. pMMO oxidizes methane and linear short-chain hydrocarbons but not aromatic compounds, the alicyclic hydrocarbon cyclohexane or the branched aliphatic 2-methylpropane, all of which are substrates of sMMO (reviewed in Smith and Dalton, 2004). Thus it seems that access to the active site of pMMO is sterically more restricted than in the soluble enzyme. Consistent with this, acetylene is a potent suicide substrate of both pMMO and sMMO, whereas the larger phenylacetylene molecule is much more effective against sMMO (Lontoh et al., 2008).

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Biochemistry and Molecular Biology of Methane Monooxygenase

Biochemistry of Soluble Methane Monooxygenase (sMMO)

sMMO is a three-component binuclear iron active centre monooxygenase that belongs to a large group of bacterial hydrocarbon oxygenases (reviewed in Leahy et al., 2003) known as the soluble diiron monooxygenases (SDIMOs) (Coleman et al., 2006), which are also homologous to the R2 subunit of class I ribonucleotide reductase. sMMO is currently the better characterized form of MMO since it is more easily purified than the particulate enzyme. More is known about the molecular mechanisms regulating expression of sMMO and a system for expression of recombinant sMMO, a prerequisite for site-directed mutagenesis studies, has also been developed (Smith et al., 2002). The most well-characterized sMMO systems are from Methylococcus capsulatus (Bath) and Methylosinus trichosporium OB3b. sMMO, encoded by a six-gene operon mmoXYBZDC has three components: (1) a 250-kDa hydroxylase with an (abg)2 structure (encoded by mmoX, Y and Z respectively). MmoX contains the binuclear iron active centre where substrate oxygenation occurs; (2) a 39-kDa NAD(P)H-dependent reductase (MmoC) with flavin adenine dinucleotide (FAD) and Fe2S2 prosthetic groups; (3) a 16-kDa component (MmoB) known as protein B or the coupling/gating protein that does not contain prosthetic groups or metal ions. The role of MmoD is still uncertain. There are X-ray crystal structures for the hydroxylase component (Elango et al., 1997; Rosenzweig et al., 1993), NMR-derived structures for protein B (Walters et al., 1999) and NMR structural data for the flavin domain of the reductase (Chatwood et al., 2004). The complex formed by the three components has been studied structurally via small angle X-ray scattering analysis and biophysically by electron paramagnetic resonance, ultracentrifugation and calorimetric analysis (reviewed in Hakemian and Rosenzweig, 2007). The catalytic cycle of sMMO has been extensively studied and excellent progress has been made towards understanding the mechanism of oxygen and hydrocarbon activation at the binuclear iron centre. More detailed reviews and descriptions of the intermediates that are known in the catalytic cycle of sMMO can be found elsewhere (Baik et al., 2003; Smith and Dalton, 2004). In order to comprehend the remarkable ability of sMMO to oxidize the unreactive methane molecule, the most noteworthy intermediate is the so-called compound Q. Compound Q accumulates when the reduced (FeII-FeII) hydroxylase is reacted with O2 in the presence of protein B. The active centre of compound Q is in a high-valent state, most likely diferryl (FeIV-FeIV). In the absence of oxidizable substrates compound Q is astonishingly stable (t1/2
14 s in aqueous solution at 4 C); however, this intermediate rapidly oxidizes methane and other substrates and is kinetically competent, i.e., the oxidation rate observed is high enough to account for the rate seen during steady state catalysis. The mechanism via which sMMO breaks the unreactive C-H bond of methane continues to be intensely debated (as reviewed in Baik et al., 2003; Hakemian and Rosenzweig, 2007; Jin and Lipscomb, 2000). Radical, ionic and concerted mechanisms have been suggested. Current evidence, based on the use of radical clock substrates and theoretical studies, suggests a reaction with multiple pathways and the possible involvement of a captive substrate-derived radical species. Recent results from the Lipscomb group have established the involvement of quantum mechanical tunneling of hydrogen nuclei in breaking the C-H bond of methane (Zheng and Lipscomb, 2006). The active site pocket of sMMO is a hydrophobic cavity deeply buried in the protein, which has been shown by molecular docking studies to be the energetically most favorable place for binding of methane and other small substrates, and clearly substrates as large as

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di-aromatics must be able to gain access to this cavity and the adjacent binuclear iron centre. The side chain of residue Leu 110 in the a-subunit of the hydroxylase partly blocks the aperture between the substrate binding pocket and the innermost of a chain of cavities that communicate between the active centre and the outside and may form the route for substrate entry and product exit. Leu 110 exhibits different conformations in different crystal forms of the enzyme. In the ‘‘closed’’ conformation it blocks access to the active site whilst in the ‘‘open’’ conformation it permits a 2.6-A˚ diameter channel into the substrate-binding cavity. A larger conformational change, which may be caused by interaction with the other components of the sMMO complex, could open this ‘‘leucine gate’’ further, to allow passage of substrates and products (Rosenzweig et al., 1997). Recent site-directed mutagenesis studies have indicated that Leu 110 is important in determining the precision with which aromatic substrates can be oriented in the active site but is not the limiting factor on the size of substrate that can enter (Borodina et al., 2007). Whilst much remains to be discovered about the molecular mechanism of substrate recognition and oxidation by sMMO, it is clear that this enzyme produces in its active site one of the most powerful oxidizing agents in nature and has a substrate-binding pocket that can accommodate a wide range of oxidation substrates in addition to the natural substrate methane.

4

Molecular Biology and Regulation of Methane Monooxygenases

In the chromosome of Mc. capsulatus Bath, there are two copies of the pMMO gene cluster pmoCAB and an additional copy of pmoC (Stolyar et al., 2001). Duplication of genes amoCAB, encoding the ammonia monooxygenase (AMO) in nitrifying bacteria has also been observed and it has been suggested that both pMMO and AMO enzymes may be evolutionarily related. A high degree of homology of pMMOs (80–94%) and duplication of pmoCAB genes also occurs in the Type II methanotrophs Ms. trichosporium and Methylocystis. Recently in type II methanotrophs two very different pmoA genes were found: conventional pmoA or pmoA1 and novel pmoA or pmoA2 (Tchawa Yimga et al., 2003). In Methylocystis strain SC2 pmoA1 and pmoA2 gene copies are each part of a complete pmoCAB gene cluster (pmoCAB1 and pmoCAB2) which exhibit low levels of identity at both the DNA level (67.4–70.9%) and the derived protein level (59.3–65.6%) but the secondary structures predicted for PmoCAB1 and PmoCAB2, as well as the derived transmembrane-spanning regions, are nearly identical (Ricke et al., 2004). Recently it has been shown that the conventional pMMO genes encode a pMMO that is expressed and oxidizes methane only at high concentrations (>600 ppmv) whereas pmoCAB2 encoding the more unusual isoenzyme pMMO2 is constitutively expressed and oxidizes methane at low concentrations, even at the trace levels of atmospheric methane (2 ppmv) (Baani and Liesack, 2008). This may well be the MMO enzyme system present in soils which for a long time have been observed to be dominated by type II methanotrophs and which oxidize methane at atmospheric concentrations. In Mc. capsulatus Bath, six ORFs organized in one operon mmoXYBZDC encode the structural genes for sMMO. The exact mechanism of reciprocal regulation of sMMO and pMMO synthesis by Cu ions is not known. Transcription of the mmo operon is initiated from a sn-(s54)-dependent promoter which requires a transcriptional activator for the formation of an active transcriptional complex. Located near the structural genes in the sMMO gene

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Biochemistry and Molecular Biology of Methane Monooxygenase

cluster of Mc. capsulatus Bath and Ms. trichosporium OB3b are two additional genes mmoR and mmoG. MmoR encoded by mmoR belongs to a class of transcriptional activators which enhance binding of RNA polymerase sN (RpoN) to promoters which are regulated by this alternative s factor. MmoG is a homologue of the chaperonin GroEL, and may be required for assembly of MmoR or indeed for assembly of the sMMO complex itself. (Czaki et al., 2003; Stafford et al., 2003). Mutagenesis of mmoR, mmoG or rpoN in these methanotrophs prevents expression of sMMO. Recently two copies of mmoX have been observed in Methylosinus sporium 5, however, mutagenesis of the second copy of mmoX which occurs on its own in the chromosome and is separate from the usual mmoXYBZDC cluster showed that this second copy is not functional. During growth of methanotrophs that contain both pMMO and sMMO under conditions where there is a low copper-to-biomass ratio, transcription of mmoR and mmoG and correct folding of MmoR may occur. The latter may then form a complex with RNA polymerase containing sN which facilitates transcription of mmoXYBZDC. Alternatively, during growth in medium where there is a high copper-to-biomass ratio, MmoR is inactivated directly or via MmoG by an as yet unknown mechanism. Two further genes, mmoQ and mmoS, which are homologous to two-component signaling systems in other bacteria, are found adjacent to the structural and regulatory genes in Mc. capsulatus (Bath) and could be involved in copper sensing. However, the exact mechanisms by which copper interacts directly (or indirectly) with MmoR to prevent transcription, or how the cells sense the intracellular or extracellular levels of copper which switch of expression of sMMO, are unclear (reviewed in Hakemian and Rosenzweig, 2007). Expression of the pmoCAB cluster during growth on medium containing excess copper ions occurs via a s70 activated promoter located 50 of pmoC. In the absence of copper ions, pMMO genes are still expressed, albeit at lower levels but the apo-enzyme produced is inactive. This inactive pMMO can be activated in vitro by the addition of copper ions. Again the exact mechanism by which pMMO is regulated is not known. Interestingly in Methylocella silvestris, which does not contain pMMO, the expression of soluble MMO is not repressed by copper ions but instead is repressed by the presence of multi-carbon substrates such as acetate. Methanobactin, a copper-chelating siderophore-like molecule of 1,217 Da binds copper with high affinity. Methanobactin was first isolated from spent medium of Ms. trichosporium and Mc. capsulatus grown at low copper and the metal binding properties of this chalkophore have been studied in some detail (e.g., see Choi et al., 2005, 2006; DiSpirito et al., 1998; Kim et al., 2004, 2005). Its crystal structure has also been elucidated. Methanobactin is probably involved in copper uptake and may also play a role in pMMO activity (reviewed in Balasubramanian and Rosenzweig, 2008).

5

Methanotrophs in Biocatalysis and Bioremediation

Interest in methanotrophic bacteria as biocatalysts for synthetic chemistry and bioremediation stems almost exclusively from the unique catalytic properties of the two MMO systems, most importantly their ability (a) to oxidize methane to methanol and (b) to co-oxidize a wide range of other substrates. Both systems require an exogenous source of reductant for the monooxygenation reaction, which in whole-cell applications can be supplied from added methanol or formate, via the principal enzymes of methylotrophic metabolism that are also present in the cells. In addition, the presence of oxygen-stable hydrogenase activity in methanotrophs enables hydrogen to be used as the reductant.

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sMMO can co-oxidize a remarkable range of alkanes, alkenes, cyclic alkanes, aromatic compounds and substituted aliphatic and aromatic compounds even though methanotrophs cannot grow on these compounds (reviewed in Smith and Dalton, 2004; Smith and Murrell, 2008). Singly oxygenated products predominate with all substrates. Alkanes are hydroxylated, in the case of aliphatic compounds almost exclusively at the terminal and subterminal positions. Ring hydroxylation of aromatics occurs primarily at the meta position, along with a comparable amount of substituent hydroxylation when an alkyl substituent is present. sMMO oxygenates alkenes to epoxides with retention of stereochemistry around the C = C double bond. Ethers are cleaved oxidatively to yield mixtures of alcohols and aldehydes, and pyridine undergoes N-oxygenation. The initial oxygenated products formed from halogenated substrates may decompose rapidly via non-enzymatic pathways that result in the loss of halogen substituents. It is certain that there are many substrates of sMMO that have simply never been tested with the enzyme. A very few small organic compounds are known not to be effective substrates of sMMO. These include tetrachloromethane, iodomethane, trimethylamine and tetrachloroethene (reviewed in Smith and Dalton, 2004; Smith and Murrell, 2008). An extensive study was performed by Dalton and co-workers toward developing sMMOexpressing Mc. capsulatus cells for production of epoxypropane from propane. In this pilot process, methanol was used as the reductant and inhibition of sMMO by the epoxide product was overcome by operating the process in a continuous two-stage system that allowed epoxide-inhibited culture to recover in a separate bioreactor in the presence of methane and other nutrients. The process gave good productivity and had the advantage that at 45 C (the optimal growth temperature of Mc. capsulatus) the epoxypropane product was easily recovered from the gas phase. With cells at 30 g L1 the epoxypropane production rate was 250 g L1 day1 and the total cost of epoxypropane production was estimated at US$1.26 per kg (Richards et al., 1994). The process came close to reaching the same cost as the established commercial chemical technology but did not offer a financial advantage over the existing technology so has not yet been commercialized, although patents for the process have been filed worldwide. The process was also evaluated for production of 1,2-epoxybutane from but1-ene and acetaldehyde from ethane. The diverse co-oxidation reactions catalyzed by sMMO and pMMO have led to many suggested applications in the oxidation of environmental pollutants (reviewed in Smith and Dalton, 2004). The priority pollutant trichloroethylene (TCE) is a substrate for both forms of MMO (see Lee et al., 2006) and, by a combination of enzyme-catalysed oxygenation and nonenzymatic steps, pMMO-expressing methanotroph cells can lead to its mineralization to CO2, water and chloride. There has been a large number of pilot studies into the use of methaneoxidizing bacteria for bioremediation of groundwater and effluents contaminated with TCE and other chlorinated solvents. During a long-term study, a TCE-contaminated aquifer in Japan has been periodically biostimulated with methane and inorganic nutrients to promote growth of methanotrophic bacteria to degrade the TCE. Here a stable and significant (10%) decrease from the initial concentration of TCE (200 ppb) was observed from 40 days after beginning biostimulation with methane. The TCE concentration returned to its initial level after biostimulation ceased. Pilot ex situ systems for bioremediation of chlorinated organic solvents using methanotrophs have included practical and financial evaluation of a two-stage process where a mixed methanotroph culture was employed at low copper-to-biomass ratio (to promote sMMO expression) in order to purify effluent contaminated with TCE and cis1,2-dichloroethylene (cDCE). Here, up to 99% removal of TCE or cDCE (initial concentration 2.25 mg L1) was achieved. Competition between methane and the chlorinated co-substrate for the (s)MMO active site was avoided by growing the cells on methane in the growth reactor

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Biochemistry and Molecular Biology of Methane Monooxygenase

. Figure 1 Principal metabolic pathways of methanotrophic metabolism showing the roles of soluble and particulate methane monooxygenase (s/pMMO), methanol dehydrogenase (MDH) and the cyclical pathways for carbon fixation. Biotechnologically important reactions and products discussed in the text are shown as bullet points.

. Figure 2 Schematic of the sMMO enzyme complex.

and then mixing with the contaminated wastewater in the second stage reactor (a plug flow reactor), where formate was added in the absence of methane to supply the reducing equivalents required by MMO (see reviews by Smith and Dalton, 2004; Smith and Murrell, 2008 and references therein for further detail on bioremediation and biocatalysis by methanotrophs).

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. Figure 3 Structure of the putative active site of sMMO, showing the substrate binding pocket (right) adjacent to the di-iron active center and cavity 2 (left), linked via the ‘‘leucine gate’’ Leu 110. The proposed route for substrate entry is indicated.

Other possibilities for bioremediation using methanotrophs include use of sMMOexpressing cells to facilitate biodegradation of mono- and di-aromatic pollutants (including polychlorinated biphenyls) by introducing oxygen functionality into these recalcitrant molecules. In the longer term, methanotrophs expressing recombinant sMMO enzymes with increased substrate range or regioselectivity may be developed for novel biotechnological applications using the mutagenesis system mentioned earlier (reviewed in Smith and Murrell, 2008).

6

Research Needs

There are a still a number of challenges in the study of methane monooxygenases and their regulation. Thus far there is no good expression system for pMMO, a prerequisite for sitedirected mutagenesis studies. This will need to be overcome before the amino-acyl residues involved in catalysis can be fully identified and the exact catalytic mechanism can be elucidated. The exact nature and function of the copper centres and the di-iron centre recently identified in pMMO need to be defined. Also the in vivo electron donor and pathways of electron transfer to pMMO are not yet known. It will also be interesting to learn the exact function of methanobactin in copper sequestration and delivery of copper ions to the active site of pMMO. The availability of the genome sequence of Mc. capsulatus, together with a facile genetic system will facilitate the study of copper transport/uptake systems in methanotrophs and help determine exactly how copper regulates the expression of sMMO and pMMO in methanotrophs that contain both enzyme systems. The role of two-component systems in methanotrophs with respect to regulation of methane oxidation also needs attention as does the mechanism of regulation of sMMO by multi-carbon compounds in Methylocella silvestris. A good system for the expression and mutagenesis of sMMO from Ms. trichosporium is now available. This will enable researchers to define the exact nature of the active site of sMMO and what makes this enzyme unique in being able to oxidize methane, and also enable mutation of sMMO in order to alter its catalytic utility. This fascinating enzyme already co-oxidizes a wide range of organic substrates and it will be interesting to ascertain if its

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properties can be further enhanced by either random or directed mutagenesis and gene shuffling, for example to co-oxidize polyaromatic hydrocarbons or make chiral epoxides or alcohols, thus improving its biotechnological potential even further.

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Biochemistry and Molecular Biology of Methane Monooxygenase the biological oxidation of methane. Nature 434: 177–182. Lontoh S, DiSpirito AA, Krema CL, Whittaker MR, Hooper AB, Semrau JD (2008) Differential inhibition in vivo of ammonia monooxygenase, soluble methane monooxygenase and membrane-associated methane monooxygenase by phenylacetylene. Env Microbiol 2: 485–494. Martinho M, Choi DW, DiSpirito AA, Antholine WE, Semrau JD, Mu¨nck E (2007) Mo¨ssbauer studies of the membrane-associated methane monooxygenase from Methylococcus capsulatus Bath: evidence for a diiron center. J Am Chem Soc 129: 15783–15785. McDonald IR, Bodrossy L, Chen Y, Murrell JC (2008) Molecular ecology techniques for the study of aerobic methanotrophs. Appl Environ Microbiol 74: 1305–1315. Murrell JC, McDonald IR, Gilbert B (2000) Regulation of expression of methane monooxygenases by copper ions. Trends Microbiol 8: 221–225. Myronova N, Kitmitto A, Collins RF, Miyaji A, Dalton H (2006) Three-dimensional structure determination of a protein supercomplex that oxidizes methane to formaldehyde in Methylococcus capsulatus (Bath). Biochemistry 45: 11905–11914. Nguyen HT, Elliott SJ, Yip JH, Chan SI (1998) The particulate methane monooxygenase from Methylococcus capsulatus (Bath) is a novel coppercontaining three-subunit enzyme. J Biol Chem 273: 7957–7978. Richards AO, Stanley SH, Suzuki M, Dalton H (1994) The biotransformation of propylene to propylene oxide by Methylococcus capsulatus (Bath). Biocatalysis 8: 253–267. Ricke P, Erkel C, Kube M, Reinhardt R, Liesack W (2004) Comparative analysis of the conventional and novel pmo (particulate methane monooxygenase) operons from Methylocystis strain SC2. Appl Environ Microbiol 70: 3055–3063. Rosenzweig AC, Brandstetter H, Whittington DA, Nordlund P, Lippard SJ, Frederick CA (1997) Crystal structures of the methane monooxygenase hydroxylase from Methylococcus capsulatus (Bath): implications for substrate gating and component interactions. Proteins 29: 141–152. Rosenzweig AC, Frederick CA, Lippard SJ, Nordlund P (1993) Crystal structure of a bacterial non-haem iron hydroxylase that catalyses the biological oxidation of methane. Nature 366: 537–543. Smith DDS, Dalton H (1989) Solubilization of methane monooxygenase from Methylococcus capsulatus (Bath). Eur J Biochem 182: 667–671.

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Smith TJ, Dalton H (2004) Biocatalysis by methane monooxygenase and its implications for the petroleum industry. Petroleum Biotechnology: Developments and Perspectives Studies in Surface Science and Catalysis 151: 177–192. Smith TJ, Murrell JC (2008) Methanotrophs. In: Encyclopedia of Industrial Biotechnology. M Flickinger, (ed.). Wiley, in press. Smith TJ, Slade SE, Burton NP, Murrell JC, Dalton H (2002) An improved system for protein engineering of the hydroxylase component of soluble methane monooxygenase. Appl Environ Microbiol 68: 5265–5273. Stafford GP, Scanlan J, McDonald IR, Murrell JC (2003) Characterization of rpoN, mmoR and mmoG, genes involved in regulating the expression of soluble methane monooxygenase in Methylosinus trichosporium OB3b Microbiol 149: 1771–1784. Stanley SH, Prior SD, Leak DJ, Dalton H (1983) Copper stress underlies the fundamental change in intracellular location of methane monooxygenase in methane-oxidizing organisms: studies in batch and continuous cultures. Biotechnol Lett 5: 487–492. Stolyar S, Franke M, Lidstrom ME (2001) Expression of individual copies of Methylococcus capsulatus Bath particulate methane monooxygenase genes. J Bacteriol 183: 1810–1812. Tchawa Yimga M, Dunfield PF, Ricke P, Heyer J, Liesack W (2003) Wide distribution of a novel pmoA-like gene copy among type II methanotrophs, and its expression in Methylocystis strain SC2. Appl Environ Microbiol 69: 5593–5602. Theisen AR, Ali HM, Radajewski S., Dumont MG, Dunfield PF, McDonald IR, Dedysh SN, Miguez CB, Murrell JC (2005) Regulation of methane oxidation in the facultative methanotroph Methylocella silvestris BL2. Mol Microbiol 58: 682–692. Trotsenko YA, Murrell JC (2008) Metabolic aspects of aerobic obligate methylotrophy. Adv App Microbiol 63: 183–229. Walters KJ, Gassner GT, Lippard SJ, Wagner G (1999) Structure of the soluble methane monooxygenase regulatory protein B. Proc Natl Acad Sci USA 96: 7877–7882. Zahn JA, DiSpirito AA (1996) Membrane-associated methane monooxygenase from Methylococcus capsulatus (Bath) J Bacteriol 178: 1018–1029. Zheng H, Lipscomb JD (2006) Regulation of methane monooxygenase catalysis based on size exclusion and quantum tunneling. Biochemistry 45: 1685–1692.

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18 Aerobic Degradation of Aromatic Hydrocarbons: Enzyme Structures and Catalytic Mechanisms J. D. Haddock Department of Microbiology, Southern Illinois University, Carbondale, IL, USA [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1058

2

Ring-Hydroxylating Dioxygenases (EC 1.14.12._) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1058

3

Cis-Dihydrodiol Dehydrogenases (EC 1.3.1._) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1061

4 Ring-Cleavage Dioxygenases (EC 1.13.11._) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1062 4.1 Intradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1062 4.2 Extradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1063 5

Bacterial Multicomponent Monooxygenases (EC 1.14.13._) . . . . . . . . . . . . . . . . . . . . . 1065

6

Xylene Monooxygenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1067

7

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1067

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_74, # Springer-Verlag Berlin Heidelberg, 2010

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Aerobic Degradation of Aromatic Hydrocarbons

Abstract: Aerobic microbial degradation pathways for aromatic hydrocarbons utilize enzymes that catalyze reactions between molecular oxygen and the substrate. Hydroxylases initiate degradation of inert aromatic hydrocarbons by addition of oxygen atoms of molecular oxygen as hydroxyl groups to the aromatic ring. Monooxygenases introduce one atom of oxygen while dioxygenases introduce both. Ring-cleavage dioxygenases decyclize aromatic compounds allowing the products to be channeled into the cell’s central metabolic pathways and support growth. Aromatic hydrocarbon oxygenases are often composed of several different protein components and subunits, but most utilize iron at the active site to bind molecular oxygen and activate it or the substrate during catalysis.

1

Introduction

Aromatic hydrocarbons are an abundant source of carbon and energy in the environment for growth of microorganisms. In order to utilize them as growth substrates, an organism must first convert these unsaturated, resonance-stabilized cyclic compounds to intermediates that can enter the central metabolic pathways. However, the chemical stability of aromatic hydrocarbons makes it difficult for microorganisms to metabolize them unless they possess specialized biodegradative pathways involving enzymes that catalyze chemical reactions that activate the aromatic ring so that it can be cleaved into fragments that are more easily metabolized. Many aerobic bacteria found in environmental habitats where aromatic compounds occur utilize molecular oxygen as a substrate to attack the aromatic ring. Mono- and dioxygenases are enzymes that catalyze hydroxylation reactions that activate aromatic rings by addition of the atoms of molecular oxygen to one or more carbon atoms of the substrate (Aerobic degradation of aromatic hydrocarbons: enzyme structures and catalytic mechanisms, (> Fig. 1). Cyclic aromatic products containing two hydroxyl substituents on adjacent carbon atoms of the ring are then converted to noncyclic products by ring-cleavage dioxygenases that cleave the C–C bond either between the hydroxylated carbons (ortho cleavage) or between a hydroxylated and nonhydroxylated carbon (meta cleavage). The pathways for degradation of diverse aromatic compounds often converge by channeling the intermediates towards production of a few common hydroxylated ring-cleavage substrates such as (methyl)catechols and protocatechuate. Following ring cleavage, subsequent reactions then yield products such as acetate, pyruvate and succinate that enter the Krebs Tricarboxylic Acid cycle or are used for biosynthesis (Fuchs, 1999). In this chapter, a few representative enzymes involved in activation and ring cleavage of aromatic hydrocarbons containing only carbon and hydrogen are examined. Analogous pathways and enzymes are involved in aerobic degradation of substituted aromatic compounds such as aromatic acids and alcohols as well as nitro-, amino-, sulfo- and chloro-substitued aromatic compounds as well as alkanes, alicyclic hydrocarbons, terpenes and fatty acids (Haddock, 2002). Those enzymes are not included in this discussion because of limited space, but descriptions of their roles and properties can be found at the University of Minnesota Biocatalysis/Biodegradation Database http://umbbd.msi.umn.edu/index.html.

2

Ring-Hydroxylating Dioxygenases (EC 1.14.12._)

Ring-hydroxylating dioxygenases oxidize aromatic rings by addition of both atoms of molecular oxygen to two adjacent carbon atoms. Two electrons are also required and the reaction

Aerobic Degradation of Aromatic Hydrocarbons

18

. Figure 1 Chemical structures of representative aromatic hydrocarbons and degradation intermediates. (a) Examples of commonly occurring aromatic hydrocarbons that are substrates for mono- and dioxygenase catalyzed reactions. I. Mono- and bicyclic aromatic hydrocarbons. R = : H, benzene; CH3, toluene; C6H5, biphenyl. II. Polynuclear aromatic hydrocarbons (PAH), naphthalene and phenanthrene, consisting of two and four fused rings, respectively. (b) Common ring-cleavage substrates produced by mono- and dioxygenase catalyzed reactions of aerobic aromatic hydrocarbon degradation pathways. R1 = : H, catechol; CH3, 3-methyl catechol; C6H5, 2,3-dihydroxy biphenyl. R2 = : CH3, 4-methyl catechol; COO
, protocatechuate. (c) Common ring-cleavage products of reactions catalyzed by extradiol (meta) and intradiol (ortho) ring-cleavage dioxygenases. R1 = : H, 2-hydroxy-6-oxo-hexa-2,4-dienoate; CH3, 2-hydroxy-6-oxo-hepta-2,4-dienoate; C6H5, 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoate. R2 = : CH3, 2-hydroxy-4-methyl-6-oxo-hexa-2,4-dienoate; COO
, 2-hydroxy-4-carboxy-6-oxohexa-2,4-dienoate. R3 = H, cis, cis-muconate. R4 = COO
, 3-carboxy-cis, cis-muconate.

products are called cis-dihydrodiols because both hydroxyl groups are located on the same side of the plane formed by the ring carbons. Toluene, naphthalene and biphenyl dioxygenase are well-characterized enzymes that are very similar in size, composition and molecular structure. They are multicomponent enzymes consisting of three separate proteins that are soluble and not membrane bound. The active site is contained in the hydoxylase component while a reductase and ferredoxin form a short electron transport chain that supplies electrons to the hydroxylase. The reductase is a flavoprotein containing FAD as a redox center that oxidizes NADH and transfers the electrons to the ferredoxin, a low molecular weight protein that

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Aerobic Degradation of Aromatic Hydrocarbons

contains a Rieske-type [2Fe–2S] iron-sulfur redox center. The iron ions of [2Fe–2S] Rieske proteins have two cysteine and two histidine ligands as opposed to plant-type ferredoxins which have four cysteine ligands (Cammack, 1992). The reduced ferredoxin transfers the electrons to the hydroxylase which contains binding sites for molecular oxygen and the aromatic substrate. The reductase of naphthalene dioxygenase is somewhat different from other dioxygenases in that it contains a plant-type iron-sulfur center in addition to FAD. The catalytic hydroxylase component consists of large (a) and small (b) protein subunits tightly associated with one another in an a3b3 quaternary structure. The crystal structure of the hydroxylase of naphthalene dioxygenase from Pseudomonas sp. strain NCIB 9816–4 reveals that the six subunits interact to produce a structure described as resembling a mushroom with the b subunits forming the stem and the a subunits forming the cap (Kauppi et al., 1998). The a subunit contains a Rieske [2Fe–2S] redox center that accepts electrons from the ferredoxin component and transfers them to a non-heme, mononuclear iron center, also located in the a subunit (Aerobic degradation of aromatic hydrocarbons: enzyme structures and catalytic mechanisms, > Fig. 2). One aspartate and two histidines are ligands to the mononuclear ferrous iron which binds and activates molecular oxygen prior to addition of the oxygen atoms to two adjacent carbon atoms in aromatic hydrocarbon substrates. The substrate-binding site is located in close proximity to the mononuclear iron-binding site. The b subunit is hypothesized to be involved in maintaining structure and/or protein-protein interaction with the ferredoxin. Crystal structures of biphenyl dioxygenase of Sphingobium yanoikuyae B1 and the ring-hydroxylating dioxygenase of Sphingomonas sp. strain CHY-1 are

. Figure 2 Proposed catalytic mechanism of aromatic hydrocarbon ring-hydroxylating dioxygenases. Some of the proposed intermediates produced during active-site binding and insertion of molecular oxygen into the ring. I. Resting state of the ferrous mononuclear iron at the active site of the hydroxylase component; II active site iron after binding of the aromatic substrate and molecular oxygen and transfer of one electron from the Rieske center; III. bound cis-dihydrodiol produced by addition of both oxygen atoms to the ring (Kovaleva and Lipscomb, 2008).

Aerobic Degradation of Aromatic Hydrocarbons

18

very similar to that of naphthalene dioxygenase. These enzymes are able to oxidize polynuclear aromatic hydrocarbons containing up to five fused aromatic rings (Ferraro et al., 2007; Jakoncic et al., 2007). The catalytic mechanism of dihydroxylation of aromatic compounds is not known in detail. Current knowledge is largely based on studies of naphthalene, benzoate and phthalate dioxygenases (Kovaleva et al., 2007). Molecular oxygen does not readily react with organic compounds and catalysis involves activation of the oxygen atoms prior to attack of aromatic substrates. Oxygen activation involves binding of O2 to the mononuclear iron of the oxygenase component and formation of an intermediate that attacks adjacent carbons of the aromatic substrate forming a cis-dihydrodiol product (> Fig. 2). Two electrons are consumed during catalysis and are supplied by the reductase and ferredoxin. Initially, one Rieske iron is Fe(III) while the other and the mononuclear iron are Fe(II). Proposed steps of the reaction involve: (1) binding of the aromatic substrate to the hydroxylase at a site that is in close proximity to the mononuclear iron, with no change in iron oxidation states; (2) binding of O2 to the mononuclear iron and transfer of one electron from the Rieske center to the mononuclear iron to form an Fe(III) hydroperoxo intermediate; (3) O–O bond cleavage and attack of two adjacent carbon atoms of the aromatic substrate, (4) transfer of one electron from the ferredoxin component which reduces the mononuclear iron to Fe(II) and breaks the Fe–O bonds to release the dihydrodiol product; (5) finally, a second electron from ferredoxin is transferred to one of the Rieske irons reducing it to Fe(II) to return the hydroxylase to the initial state. Alternatively, at step 2/3, the Fe(III) hydroperoxo intermediate may be converted to an Fe(V)-oxo-hydrooxo species which attacks the aromatic substrate.

3

Cis-Dihydrodiol Dehydrogenases (EC 1.3.1._)

cis-Dihydrodiol dehydrogenases are enzymes of aerobic aromatic compound degradation pathways that catalyze oxidation and aromatization of cis-dihydrodiols produced by ringhydroxylating dioxygenases. The resulting catechols are then substrates for ring-cleavage dioxygenases. The dehydrogenases are usually a4 homotetramers with a native size of approximately 110,000 kDa. They reduce NAD+ to NADH when catalyzing oxidation of cisdihydrodiols (Jouanneau and Meyer, 2006). The crystal structure of cis-2,3-dihydroxy-2,3dihydrobiphenyl dehydrogenase of Burkholderia xenovorans LB400 with bound NAD+ has been reported (Hulsmeyer et al., 1998). The 277 amino acid monomer is a member of the short-chain alcohol dehydrogenase/reductase (SDR) enzyme family. The active site contains a Ser-Tyr-Lys triad that is conserved in SDR enzymes. The substrate binds in a deep hydrophobic cleft with the hydroxylated ring near the nicotinamide ring of bound NAD+. A tryptophan residue may facilitate catalysis by affecting the dihedral angle between the two rings of the bicyclic substrate. A conserved asparagine residue may be involved in determining substrate specificity by hydrogen bonding to one of the substrates hydroxyl groups. This amino acid is replaced by valine in the naphthalene pathway enzyme and threonine in Sphingomonas sp. strain CHY-1 (Jouanneau and Meyer, 2006). Oxidation of the biphenyl dihydrodiol intermediate involves transfer of a hydride ion from the C3 position to NAD+. In the first step of the reaction, a deprotonated tyrosine residue is thought to remove the proton from the C3 hydroxyl group allowing the C3 hydrogen to be transferred to NAD+ yielding a cyclic hydroxy-2,4-dienone as the immediate product which then rearranges via keto-enol tautomerization to aromatize the ring.

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Aerobic Degradation of Aromatic Hydrocarbons

Sphingomonas sp. strain CHY-1 utilizes a dioxygenase pathway to grow on polynuclear aromatic hydrocarbons containing two (naphthalene), three (anthracene and phenanthrene) and four (chrysene) fused rings (Demaneche et al., 2004). The dioxygenase and its companion cis-dihydrodiol dehydrogenase have broad substrate specificity for substrates containing 2–5 fused rings. The dehydrogenase is very similar to the biphenyl dihydrodiol dehydrogenase described above. However, the conserved asparagine potentially involved in determining substrate specificity is replaced by threonine in CHY-1 and other sphingomonads (Jouanneau and Meyer, 2006).

4

Ring-Cleavage Dioxygenases (EC 1.13.11._)

Ring-cleavage dioxygenases are enzymes that catalyze a reaction in which molecular oxygen is inserted into a C–C bond of dihydroxylated aromatic compounds, resulting in cleavage of the ring. Their substrates are central intermediates of aerobic aromatic compound degradation pathways. The ring-cleavage products are then further metabolized to compounds that enter the central metabolic pathways, allowing the microorganisms to obtain energy and carbon substrates for biosynthesis. Examples of substrates and products are shown in > Fig. 1. The enzymes are distinctly different from the ring-hydroxylating dioxygenases described above. Ring-cleavage dioxygenases can be placed into two major classes, intradiol or extradiol, depending on the position of cleavage relative to that of the ring hydroxyl groups. Vaillancourt et al. (2006) have composed a thorough summary of the extensive research findings by themselves and others on the ring-cleavage dioxygenases.

4.1

Intradiol Dioxygenases

Intradiol ring-cleavage dioxygenases utilize non-heme Fe(III) as a cofactor to catalyze insertion of both atoms of molecular oxygen into catechols to break the bond between two adjacent ring carbon atoms bearing hydroxyl groups (ortho cleavage). The ortho-cleavage product of catechol is cis,cis-muconate (> Fig. 1). A broad diversity of bacterial species utilizes these enzymes but all of the proteins appear to be evolutionarily related and have similar properties. The main differences are in subunit composition and quaternary structure. Catechol 1,2-dioxygenase (C–12O) of Acinetobacter baylyi ADP1 (formerly A. calcoaceticus ADP1) is an a2 homodimer (34,342 kDa/monomer). Each subunit contains a catalytic domain containing an Fe(III) ion bound by two tyrosine and two histidine ligands with hydroxide as a fifth ligand. A dimerization domain is responsible for contacts between subunits (Vetting and Ohlendorf, 2000). Protocatechuate 3,4-dioxygenases purified from various bacteria contain a and b subunits having configurations ranging from (ab)2 to (ab)12. The crystal structure of the enzyme from strain ADP1 shows an (abFe3+)12 quaternary structure (Vetting et al., 2000). The a and b subunits are similar in size and structure but the two tyrosine and two histidine Fe(III) ligands are located in the b subunit. The catalytic mechanism of intradiol dioxygenases is thought to involve activation of the substrate for electrophilic attack by O2 (Vaillancourt et al., 2006). The initial state of the enzyme is with the Fe(III) coordinated to two histidines, two tyrosines and hydroxide. The enzyme employs an ordered mechanism with the substrate binding first as a dianion to Fe(III), displacing one tyrosine and the hydroxide (Aerobic degradation of aromatic

Aerobic Degradation of Aromatic Hydrocarbons

18

hydrocarbons: enzyme structures and catalytic mechanisms, > Fig. 3). A substrate radical is formed by transfer of one electron to iron followed by binding of molecular oxygen which then attacks the substrate producing an iron-alkylperoxo intermediate similar to that of extradiol dioxygenases (see below). Criegee rearrangement and O–O bond cleavage result in formation of a cyclic anhydride containing one O atom with the other bound to Fe(III) as hydroxide. The hydroxide then hydrolyzes the anhydride, cleaving the bond between the carbon atoms that carried the hydroxyl groups of the substrate. The ring-cleavage product bearing two terminal carboxyl groups is then released from the enzyme.

4.2

Extradiol Dioxygenases

Extradiol ring-cleavage dioxygenases utilize non-heme Fe(II) as a cofactor to catalyze insertion of both atoms of molecular oxygen into catechols to break the bond between vicinal hydroxylated and nonhydroxylated carbons (meta cleavage). The meta-cleavage product of catechol is 2-hydroxy-6-oxo-hexa-2,4-dienoate, also called 2-hydroxy-cis,cis-muconic semialdehyde (> Fig. 1). As with the intradiol enzymes, extradiol dioxygenases are also found in a broad diversity of bacterial species. However, the two types of enzymes are evolutionary distinct from one another and extradiol dioxygenases utilize non-heme Fe(II) as a cofactor. Structurally,

. Figure 3 Proposed mechanism of aromatic hydrocarbon intradiol ring-cleavage dioxygenases. Some of the proposed intermediates formed during active-site binding and insertion of molecular oxygen into the ring. I. Ferric active-site iron bound to dianionic catechol; II. activation of the substrate; III. attack of the substrate by bound molecular oxygen; IV. intradiol ring-cleavage product prior to release from the active site (Borowski and Siegbahn, 2006; Pau et al., 2007).

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Aerobic Degradation of Aromatic Hydrocarbons

many of the extradiol enzymes consist of one type of subunit with an a2 to a8 quaternary structure. The Sphingomonas paucimobilis SYK6 protocatechuate 4,5-dioxygenase is an a2b2 heterotetramer and the Commamonas testosteroni T-2 enzyme is an a4b4 heterooctomer (Vaillancourt et al., 2006). Type I extradiol dioxygenases consist of one subunit, but may possess one or two domains. One-domain subunits are around 21 kDa in size. Two-domain enzymes are about 32.5 kDa and consist of a catalytically inactive N-domain and a C-domain containing the active site with bound Fe(II). 2,3-Dihydroxybiphenyl 1,2-dioxygenase (DHBDLB400) from B. xenovorans LB400 and Pseudomonas sp. strain KKS102 is a two domain a8 homohexamer (Hans et al., 1995; Senda et al., 1996). The active site is a funnel that is open at both ends. The larger opening is proposed to allow entry of the substrate while O2 and water may enter at either end. Five ligands to Fe(II) include two histidines, one glutamate and two water molecules. The catalytic mechanism of extradiol dioxygenases involves activation of molecular oxygen for nucleophilic attack on the substrate (Kovaleva et al., 2007; Siegbahn and Haeffner, 2004; Vaillancourt et al., 2006). The ordered mechanism involves binding of catechol substrates followed by binding of molecular oxygen (Aerobic degradation of aromatic hydrocarbons: enzyme structures and catalytic mechanisms, > Fig. 4). Catechols bind to Fe(II) as their respective monoanion with the two oxygen atoms coordinated to the iron. The bound

. Figure 4 Proposed catalytic mechanism of aromatic hydrocarbon extradiol ring-cleavage dioxygenases. Some of the proposed intermediates formed during active-site binding and insertion of molecular oxygen into the ring. I. Ferrous active-site iron bound to monoanionic catechol; II. activation of bound molecular oxygen and substrate; III. attack of the substrate by molecular oxygen; IV. extradiol ring-cleavage product prior to release from the active site (Kovaleva et al., 2007; Shu et al., 1997; Vaillancourt et al., 2002).

Aerobic Degradation of Aromatic Hydrocarbons

18

substrate activates the iron for binding to molecular oxygen producing an iron-alkylperoxo intermediate. Next, the O–O bond is cleaved and one oxygen atom is inserted into the ring producing a lactone. Ring cleavage is completed when the second oxygen atom, bound to iron as a hydroxide ion, is inserted into the lactone in a hydrolysis reaction. The ring-cleavage product, containing a carboxyl group on one end and an aldehyde on the other (i.e., a semialdehyde), is then released from the enzyme.

5

Bacterial Multicomponent Monooxygenases (EC 1.14.13._)

Bacterial multicomponent monooxygenases (BMMs) acting on aromatic hydrocarbons catalyze addition of one oxygen atom of O2 to the ring or a methyl substituent of aromatic hydrocarbons such as toluene. The other oxygen atom is reduced to water. The hydroxylated aromatic products are substrates for the same or a different enzyme that catalyzes a second monooxygenation reaction to produce catechols that can be acted on by the ring-cleavage dioxygenases described above. Aromatic hydrocarbon BMMs are members of an enzyme family that includes soluble methane monooxygenase (sMMO) and toluene monooxygenases (TMOs) (Sazinsky et al., 2004). The enzymes are composed of a hydroxylase component with an active site containing a diiron center bound to the protein by four glutamate and two histidine residues, a [2Fe–2S]-NADH-oxidizing flavoprotein reductase and a regulatory protein. The diiron center of the hydroxylase binds and activates molecular oxygen for attack of the substrate. The BMM family has been subdivided into 5–6 groups (Notomista et al., 2003). Groups 1 (phenol hydroxylases) and 2 (toluene-benzene monooxygenases) catalyze hydroxylation of aromatic hydrocarbons. The catalytic component of these enzymes is a multisubunit hydroxylase composed of three proteins arranged as a2b2g2 heterohexamers in association with a regulatory protein and a NADH oxidoreductase that supplies electrons to the hydroxylase during catalysis. The diiron center is located in the a subunit. The regulatory protein, also called the effector, is not required for activity but increases the reaction rate. Group 2 monooxygenases contain an additional electron transport component; a Rieske-type ferredoxin that transfers electrons from the reductase to the hydroxylase. Several aerobic toluene degradation pathways employ BMMs to initiate attack of the substrate (Parales et al., 2008). Toluene 2-monooxygenase (T2MO) of Burkholderia vietnamiensis G4 is a Group 1 BMM that catalyzes monooxygenation of C2 and then C3 of toluene to produce 3-methylcatechol. The enzyme is composed of three components: a 211-kDa hydroxylase, a 40-kDa reductase and a small 10.5 kDa effector protein. The hydroxylase consists of two molecules each of 54.5, 37.7 and 13.5 kDa proteins (i.e., a2b2g2). One diiron center is located within each a subunit and is involved in catalyzing monohydroxylation of toluene at C2 and then o-cresol at C3. The product, 3-methyl catechol, is a substrate for a meta-ring-cleavage dioxygenase. The reductase contains FAD and a [2Fe–2S] center that catalyzes oxidation and transfer of electrons to the hydroxylase. The effector protein is not required but increases the reaction rate approximately 10X. Phenol hydroxylases of Pseudomonas sp. strain CF600 and Pseudomonas stutzeri OX1 are also members of Group 1 BMMs (Cafaro et al., 2004). Toluene 3-monooxygenase (T3MO) of Ralstonia pickettii PK01 catalyzes hydroxylation of C3 of toluene to produce m-cresol and C2 of m-cresol to produce 3-methyl catechol, the same meta-cleavage dioxygenase substrate produced in the T2MO pathway. However it is a Group 2 BMM that utilizes a Rieske-type ferredoxin to transfer electrons from the reductase to the hydroxylase. The enzyme is also capable of catalyzing hydroxylation of C4, producing

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Aerobic Degradation of Aromatic Hydrocarbons

p-cresol, but the significance of this activity in the degradation pathway is unclear (Parales et al., 2008). Toluene 4-monooxygenase (T4MO) of Pseudomonas mendocina KR1 catalyzes hydroxylation of C4 of toluene to produce p-cresol. It is a Group 2 BMM consisting of four components as described above for T3MO. However, the enzyme participates in a toluene degradation pathway that involves oxidation of the methyl group of p-cresol and orthoring cleavage of protocatechuate to produce 3-carboxy-cis, cis-muconate (Whited and Gibson, 1991). Toluene/o-xylene monooxygenase (ToMO) of Pseudomonas stutzeri OX1 is a Group 2 BMM that exhibits relaxed regiospecificity for the position of substrate hydroxylation producing o-, m- and p-cresol. The enzyme also catalyzes hydroxylation of benzene and several other aromatic hydrocarbons (Bertoni et al., 1996). The catalytic mechanism employed by BMMs involves binding and activation of molecular oxygen to a diiron center at the active site of the hydroxylase component (Schwartz et al., 2008). The structure of the active site of TMOs appears to be highly homologous to that of sMMO (Sazinsky et al., 2004). The diiron center of both types of enzyme is bound to the enzyme by four glutamate and two histidine residues, with one glutamate providing a carboxylate bridge between the iron ions (Aerobic degradation of aromatic hydrocarbons: enzyme structures and catalytic mechanisms, > Fig. 5). The catalytic mechanism of TMOs is not as well understood as that of sMMO. The iron ions of ToMO are initially in the Fe(III)

. Figure 5 Proposed catalytic mechanism of aromatic hydrocarbon BMMs. Some of the proposed intermediates formed during active site binding and insertion of one atom of molecular oxygen into the ring. I. Binuclear Fe(III) core at the active of the hydroxylase a subunit; II. after reduction to a Fe(II)2 intermediate; III. after binding of molecular oxygen to form a peroxo diiron intermediate; III. ! I. monohydroxylation of an aromatic hydrocarbon substrate (R = CH3, toluene) (Kovaleva et al., 2007; Murray and Lippard, 2007; Shu et al., 1997).

Aerobic Degradation of Aromatic Hydrocarbons

18

oxidation state. During catalysis, they are reduced to Fe(II), followed by binding of molecular oxygen to produce a peroxo diiron (III) intermediate which is thought to monohydroxylate the substrate ring. Higher-valent oxidation states (e.g., Fe(IV)) of the diiron center observed during the catalytic cycle of sMMO have not been detected (Murray et al., 2007).

6

Xylene Monooxygenase

Xylene monooxygenase catalyzes hydroxylation of the methyl group of toluene and xylenes in a toluene degradation pathway in Pseudomonas putida mt-2. (Suzuki et al., 1991). One atom of molecular oxygen is added to toluene producing benzyl alcohol and the other atom is reduced to water. Other pathway enzymes then catalyze oxidation to benzoate, dihydroxylation and meta-ring cleavage. Xylene monooxygenase is composed of two components. XylM is a 42kDa catalytic component located in the membrane that contains a diiron center at the active site. The two iron ions are hypothesized to be bound to the protein by histidine ligands. XylA is a 40-kDa flavoprotein reductase with a plant-type [2Fe–2S] center. The enzyme has been difficult to study because of its association with the cell membrane and the catalytic mechanism and three-dimensional structure are unknown. However, XylM is similar to the AlkB hydroxylase of microorganisms that utilize alkanes for growth (Austin et al., 2003; van Beilen, et al. 2005).

7

Research Needs

Oxygenase enzymes involved in aerobic degradation pathways of aromatic hydrocarbons have been investigated since the middle of the past century (Hayaishi, 1974). Recent advances in knowledge of these enzymes have occurred through the application of sophisticated spectroscopic methods, comparison of genomes, genes and amino acid sequences from diverse microorganisms and elucidation of molecular structures. Further advances will come from continued application of these methods to expand studies to a greater diversity of microorganisms present in environments where aromatic hydrocarbons are abundant. Application of the knowledge gained may be employed in the design and modification of enzymes and pathways for use as biocatalysts to produce useful organic compounds and for biodegradation of environmental pollutants.

References Austin RN, Buzzi K, Zylstra GJ, Groves JT (2003) Xylene monooxygenase, a membrane-spanning non-heme diiron enzyme that hydroxylates hydrocarbons via a substrate radical intermediate. J Biol Inorg Chem 8: 733–740. Bertoni G, Bolognese F, Galli E, Barbieri P (1996) Cloning of the genes for and characterization of the early stages of toluene and o-xylene catabolism in Pseudomonas stutzeri OX1. Appl Environ Microbiol 62: 3704–3711.

Borowski T, Siegbahn PEM (2006) Mechanism for catechol ring cleavage by non-heme iron intradiol dioxygenases. J Am Chem Soc 128: 12941–12953. Cafaro V, Izzo V, Scognamiglio R, Notomista E, Capasso P, Casbarra A, Pucci P, Di Donato A (2004) Phenol hydroxylase and toluene/o-xylene monooxygenase from Pseudomonas stutzeri OX1: interplay between two enzymes. Appl Environ Microbiol 70: 2211–2219.

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Cammack R (1992) Iron-sulfur clusters in enzymes: themes and variations. Adv Inorganic Chem 38: 281–322. Demaneche S, Meyer C, Micoud J, Louwagie M, Willison JC, Jouanneau Y (2004) Identification and functional analysis of two aromatic-ringhydroxylating dioxygenases from a Sphingomonas strain that degrades various polycyclic aromatic hydrocarbons. Appl Environ Microbiol 70: 6714–6725. Ferraro DJ, Brown EN, Yu C-L, Parales RE, Gibson DT, Ramaswamy S (2007) Structural investigations of the ferredoxin and terminal oxygenase components of the biphenyl 2,3-dioxygenase from Sphingobium yanoikuyae B1. BMC Struct Biol 7: 10. Fuchs G (1999) Oxidation of organic compounds. In Biology of the Prokaryotes. JW Lengeler, G Drews, HG Schlegel, (eds.). Stuttgart: Thieme, pp 187–233. Haddock JD (2002) Oxygenase enzymes: role in biodegradation. In Encyclopedia of Environmental Microbiology. G Britton, (ed.). New York: Wiley, pp 2288–2300. Hans S, Eltis LD, Timmis KN, Muchmore SW, Bolin JT (1995) Crystal structure of the biphenyl-cleaving extradiol dioxygenase from a PCB-degrading pseudomonad. Science 270: 976–980. Hayaishi O (1974) General properties and biological functions of oxygenases. In Molecular mechanisms of oxygen activation. O Hayaishi, (ed.). New York: Academic Press, pp. 1–28. Hulsmeyer M, Hecht H-J, Niefind K, Hofer B, Eltis LD, Timmis KN, Schomburg D (1998) Crystal structure of cis-biphenyl-2,3-dihydrodiol-2,3-dehydrogenase from a PCB degrader at 2.0 A resolution. Protein Sci 7: 1286–1293. Jakoncic J, Jouanneau Y, Meyer C, Stojanoff V (2007) The crystal structure of the ring-hydroxylating dioxygenase from Sphingomonas CHY-1. FEBS J 274: 2470–2481. Jouanneau Y, Meyer C (2006) Purification and characterization of an arene cis-dihydrodiol dehydrogenase endowed with broad substrate specificity toward polycyclic aromatic hydrocarbon dihydrodiols. Appl Environ Microbiol 72: 4726–4734. Kauppi B, Lee K, Carredano E, Parales RE, Gibson DT, Eklund H, Ramaswamy S (1998) Structure of an aromatic-ring-hydroxylating dioxygenase-naphthalene dioxygenase. Structure 6: 571–586. Kovaleva EG, Lipscomb JD (2008) Versatility of biological non-heme Fe(II) centers in oxygen activation reactions. Nat Chem Biol 4: 186–193. Kovaleva EG, Neibergall MB, Chakrabarty S, Lipscomb JD (2007) Finding intermediates in the O2 activation pathways of non-heme iron oxygenases. Acc Chem Res 40: 475–483.

Murray LJ, Lippard SJ (2007) Substrate trafficking and dioxygen activation in bacterial multicomponent monooxygenases. Acc Chem Res 40: 466–474. Murray LJ, Naik SG, Ortillo DO, Garcia-Serres R, Lee JK, Huynh BH, Lippard SJ (2007) Characterization of the arene-oxidizing intermediate in tomoh as a diiron (III) species. J Am Chem Soc 129: 14500–14510. Notomista E, Lahm A, Di Donato A, Tramontano A (2003) Evolution of bacterial and archaeal multicomponent monooxygenases. J Mol Evol 56: 435–445. Parales RE, Parales JV, Pelletier DA, Ditty JL (2008) Diversity of microbial toluene degradation pathways. Adv Appl Microbiol 64: 1–73. Pau MYM, Davis MI, Orville AM, Lipscomb JD, Solomon EI (2007) Spectroscopic and electronic structure study of the enzyme-substrate complex of intradiol dioxygenases: substrate activation by a high-spin ferric non-heme iron site. J Am Chem Soc 129: 1944–1958. Sazinsky MH, Bard J, Di Donato A, Lippard SJ (2004) Crystal structure of the toluene/o-xylene monooxygenase hydroxylase from Pseudomonas stutzeri OX1. J Biol Chem 279: 30600–30610. Schwartz JK, Wei P-P, Mitchell KH, Fox BG, Solomon EI (2008) Geometric and electronic structure studies of the binuclear nonheme ferrous active site of toluene-4-monooxygenase: parallels with methane monooxygnease and insight into the role of the effector proteins in O2 activation. J Am Chem Soc 130: 7098–7109. Senda T, Sugiyama K, Narita H, Yamamoto T, Kimbara K, Fukuda M, Sato M, Yano K, Mitsui Y (1996) Threedimensional structures of free form and two substrate complexes of an extradiol ring-cleavage type dioxygenase, the bphc enzyme from Pseudomonas sp. Strain KKS102. J Mol Biol 255: 735–752. Shu L, Nesheim JC, Kauffmann K, Munck E, Lipscomb JD, Que L Jr, (1997) An Fe2IVO2 diamond core structure for the key intermediate Q of methane monooxygenase. Science 275: 515–518. Siegbahn PEM, Haeffner F (2004) Mechanism for catechol ring-cleavage by non-heme iron extradiol dioxygenases. J Am Chem Soc 126: 8919–8932. Suzuki M, Hayakawa T, Shaw JP, Rekik M, Harayama S (1991) Primary structure of xylene monooxygenase: similarities to and differences from the alkane hydroxylation system. J Bacteriol 173: 1690–1695. Vaillancourt FH, Barbosa CJ, Spiro TJ, Bolin JT, Blades MW, Turner RFB, Eltis LD (2002) Definitive evidence for monoanionic binding of 2,3-dihydroxybiphenyl to 2,3-dihydroxybiphenyl 1,2-dioxygenase from UV resonance raman spectroscopy, UV/vis absorption spectroscopy, and crystallography. J Am Chem Soc 20: 2485–2496.

Aerobic Degradation of Aromatic Hydrocarbons Vaillancourt FH, Bolin JT, Eltis LD (2006) The ins and outs of ring cleaving dioxygenases. Crit Rev Biochem Mol Biol 41: 241–267. van Beilen JB, Smits THM, Roos FF, Brunner T, Balada SB, Rothlisberger M, Witholt B (2005) Identification of an amino acid position that determines the substrate range of integral membrane alkane hydroxylases. J Bacteriol 2005: 85–91. Vetting MW, D’Argenio DA, Ornston LN, Ohlendorf DH (2000) Structure of Acinetobacter strain ADP1 protocatechuate 3,4-dioxygenase at 2.2 A resolution:

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implications for the mechanism of an intradiol dioxygenase. Biochemistry 39: 7943–7955. Vetting MW, Ohlendorf DH (2000) The 1.8 A crystal structure of catechol 1,2-dioxygenase reveals a novel hydrophobic helical zipper as a subunit linker. Structure 8: 429–440. Whited GM, Gibson DT (1991) Separation and partial characterization of the enzymes of the toluene4-monooxygenase catabolic pathway in Pseudomonas mendocina KR1. J Bacteriol 173: 3017–3020.

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19 Oxidative Inactivation of Ring-Cleavage Extradiol Dioxygenases: Mechanism and Ferredoxin-Mediated Reactivation Y. Jouanneau CEA, iRTSV, LCBM, and CNRS UMR 5249, Grenoble, France [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1072 2 Oxidative Inactivation of Extradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1072 3 A Natural Repair System Dedicated to the Reactivation of Catechol 2,3-Dioxygenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1074 4 Ferredoxin-Mediated Reactivation of Catechol Dioxygenases in Bacteria Degrading Other Aromatic Hydrocarbons . . . . . . . . . . . . . . 1076 5 Occurrence of xylT-Like Genes in Bacterial Genomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1077 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1078

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_75, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Extradiol dioxygenases are ubiquitous enzymes that catalyze ring cleavage of a wide variety of aromatic compounds. Most of these enzymes contain a ferrous ion at the active site, which is bound to the polypeptide chain through a conserved triad of two histidines and one glutamic acid. During the catalytic cycle, a catecholic substrate first binds at the active site, followed by dioxygen and the ternary complex formed yields a bound superoxide that attacks the substrate, leading eventually to ring cleavage. The active site iron remains ferrous during catalysis, except when poor substrates such as chloro- or methylcatechols are processed. In such cases, the enzyme becomes inactivated through oxidation and eventually loss of its active site iron atom. In Pseudomonas putida mt-2, catechol 2,3-dioxygenase, which is involved in toluene degradation, is inactivated by 4-methylcatechol. The enzyme is however rescued by a specific reactivation system involving a [2Fe-2S] ferredoxin encoded by xylT. The role of this ferredoxin is to reduce the ferric ion of the inactive enzyme thereby regenerating the active catalyst. Recent findings indicate that the electrons needed for the XylT-mediated reactivation are provided by XylZ, a NADH-oxidoreductase that is part of the toluate dioxygenase complex. XylT analogues present in other bacteria have been shown to have a similar role in the reactivation of catechol dioxygenases involved in the degradation of various aromatic hydrocarbons including cresols, chlorobenzene and naphthalene. The occurrence and significance of ferredoxin-mediated extradiol dioxygenae repair systems in bacteria is discussed.

1

Introduction

The cleavage of the aromatic ring is a critical step in the biodegradation of aromatic hydrocarbons by aerobic bacteria. It is catalyzed by two kinds of enzyme, the intradiol and extradiol dioxygenases, which represent two classes of phylogenetically unrelated proteins (Harayama et al., 1992). Although both types of enzymes are active on similar catecholic substrates they have different structures and catalytic mechanisms (Vaillancourt et al., 2006). Also, extradiol dioxygenases appear to be more versatile than intradiol dioxygenases as they react with a wider range of substrates and intervene in a greater variety of pathways. However, they are labile enzymes which undergo oxidative inactivation, especially in the presence of poor substrates such as alkyl- or chloro-substituted catechols. Current knowledge of the mechanism underlying extradiol dioxygenase inactivation is reviewed here, in the light of recent reports describing in detail the catalytic cycle of this type of enzyme and the process of mechanism-based enzyme inactivation. A natural repair system, preventing inactivation of catechol 2,3-dioxygenase, was first discovered in Pseudomonas putida mt-2 carrying the xyl genes responsible for the degradation of toluene and xylene (Polissi and Harayama, 1993). The repair system relies on a specific [2Fe-2S] ferredoxin encoded by xylT, which has been shown to reactivate inactivated catechol dioxygenase (Hugo et al., 1998). The properties and mode of action of this ferredoxin are presented below, as well as evidence for the involvement of a reductase in the reactivation process in vivo. The occurrence of XylT-like ferredoxins in other bacteria and their implication in other degradation pathways are discussed.

2

Oxidative Inactivation of Extradiol Dioxygenases

Extradiol dioxygenases have been classified into three different subfamilies based on protein sequence analysis (Vaillancourt et al., 2006). Here, only type I enzymes, including

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the well-characterized catechol 2,3-dioxygenase from P. putida mt-2 (C23Omt-2) (Kita et al., 1999), and the 2,3-dihydroxybiphenyl 1,2-dioxygenase from Burkholderia xenovorans LB400 (DHBD) (Han et al., 1995) will be considered. These enzymes contain a non-heme FeII ion at the active site, which is bound to the polypeptide chain through two histidines and the carboxylate of one glutamate residue. The FeII ion binds the substrate and molecular oxygen, and directly participates in the ring cleavage reaction which occurs between two carbons adjacent to the diol. On the basis of spectroscopic studies of extradiol dioxygenases, a mechanism for the cleavage reaction has been proposed (Shu et al., 1995). The catalytic cycle starts with the binding of the catecholic substrate to the ferrous ion in a bidentate manner (> Fig. 1; I). Substrate binding facilitates the formation of a ternary complex with O2, which gives rise to a semiquinone-superoxo intermediate (II) by transfer of electron density from the substrate to dioxygen mediated by FeII. The latter species is thought to attack the substrate to give an alkylperoxo intermediate (III). Then, cleavage of the O–O bond, followed by Criegee rearrangement leads to a lactone intermediate and an FeII-bound hydroxide anion (IV). Hydrolysis of the lactone and release of the reaction product complete the reaction cycle. Strong experimental support for this proposed reaction mechanism came from a recent elegant study in which three enzyme-intermediate complexes, including the superoxo (II) and the alkylperoxo (III) states, were trapped a crystallized and structurally characterized (Kovaleva and Lipscomb, 2007).

. Figure 1 Proposed mechanisms for the ring-cleavage reaction and suicide inactivation by extradiol dioxygenases. The scheme is adapted from previous work on the structure of reaction intermediates (Kovaleva and Lipscomb, 2007) and mechanism-based inactivation (Vaillancourt et al., 2002).

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Oxidative Inactivation of Ring-Cleavage Extradiol Dioxygenases

During the normal course of the catalytic reaction, there is no evidence that the iron changes redox state. However, when C23Omt-2 is incubated with alkylcatechols, enzyme inactivation occurs which primarily results from the accidental oxidation of the active site FeII to FeIII (Cerdan et al., 1994). Analysis of the 4-methylcatechol-inactivated C23Omt-2 by EPR spectroscopy revealed that the enzyme contained FeIII at the active site, but the signal obtained was distinct from that of a peroxide-treated enzyme, suggesting that a ligand molecule, possibly 4-methylcathecol or an oxidation product, remained bound to the enzyme (Hugo et al., 1998). Halocatechols such as 3-chlorocatechol also cause suicide inactivation of C23Omt-2 through a mechanism which has been a matter of debate. It has been proposed that inactivation results either from the oxidation and chelation of the active site iron by the catechol or irreversible covalent modification by an acyl chloride generated during ring cleavage (Bartels et al., 1984). A more recent study showed that DHBD became inactivated during the steady-state cleavage of the catechols, and resulted from an oxidation of the active site iron (Vaillancourt et al., 2002). The substrate was found to increase both the reactivity of the enzyme for oxygen and the rate of O2-dependent inactivation, the latter being faster with poor-substrates like 3-chlorocatechol. In that study, the authors proposed that inactivation of extradiol dioxygenases during turnover involved dissociation of the enzyme-superoxo intermediate at an early step of the catalytic cycle, resulting in the oxidation of the active site iron and release of the substrate (> Fig. 1). It has been suggested that substituted catechols promote more rapid inactivation because the substituent hinders the correct positioning of the substrate or of O2 at the active site, thus slowing down the catalytic reaction and favoring abortion of the catalytic cycle. Consistent with this idea, it was found that ortho-chlorinated dihydroxybiphenyls, which promote suicide inactivation of DHBD, bind the enzyme active site in such a position that the chlorine atom might affect the binding of O2 (Dai et al., 2002). Interestingly, a catechol dioxygenase showing high sequence similarity with C23Omt-2 was found to efficiently cleave 3-chlorocatechol (Kaschabek et al., 1998). This enzyme allowed P. putida GJ31, the strain from which it was isolated, to metabolize both toluene and chlorobenzene via the meta cleavage pathway (Mars et al., 1997). Although the structural features that makes this enzyme resistant to inactivation have not been determined, one can assume that residues involved in substrate binding play an essential role in the mechanism of resistance. Supporting this idea, the activity of the extradiol dioxygenase from Sphingomonas xenophaga BN6 towards 3-chlorocatechol was enhanced in variants of the enzyme carrying amino acid replacements in a protein region thought to be involved in substrate binding (Riegert et al., 2001). It is to note that the enzyme from strain BN6 is slightly different from the GJ31 enzyme in that the former converts 3-chlorocatechol to chloromuconic semialdehyde whereas the latter produces 2-hydroxymuconate.

3

A Natural Repair System Dedicated to the Reactivation of Catechol 2,3-Dioxygenase

In P. putida mt-2, the TOL plasmid carries a cluster of genes responsible for the complete degradation of toluene. The xyl genes are organized in two operons, one of which encodes the enzymes that convert toluene to benzoate, the other enzymes that transform benzoate into central metabolites. The latter operon contains 13 genes, including the xylE gene encoding C23Omt-2, which is immediately preceded by xylT, a gene coding for a [2Fe-2S] ferredoxin (Harayama and Rekik, 1990). A xylT-deleted mutant was found to lose the ability to grow on

Oxidative Inactivation of Ring-Cleavage Extradiol Dioxygenases

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p-xylene and p-toluate, due to the irreversible inactivation of C23Omt-2 by 4-methylcatechol (Polissi and Harayama, 1993). Hence, in preventing inactivation of the catechol dioxygenase, XylT allowed strain mt-2 to grow on para-substituted alkylbenzenes, thus extending the range of substrates utilized by the bacteria. The mechanism of action of XylT was studied after purification of the protein overproduced in E. coli (Hugo et al., 1998). XylT appeared to be a [2Fe-2S] ferredoxin, showing some similarities to plant counterparts, but also with distinctive biochemical and spectroscopic properties. As predicted from the prevalence of basic residues in its sequence, XylT has an isoelectric point of 8.27. This basic character shared by XylT analogues is a unique feature among ferredoxins and might be of critical importance for its interaction with C23Omt-2 (see below). XylT also exhibited an unusual instability upon exposure to air oxygen or upon incubation at moderate temperature, with half-lives of 69 min at 25 C in air and 70 min under argon at 37 C. This instability, which resulted in the loss of the Fe-S cluster, might reflect a high degree of exposure of the cluster to the solvent, as suggested from changes in the EPR signal of the protein in the presence of glycerol (Hugo et al., 1998). The direct role of XylT in C23Omt-2 reactivation was assessed in in vitro experiments where the purified ferredoxin was incubated with the inactivated catechol 2,3-dioxygenase and 5-deazaflavin as a source of reductant. Results showed that XylT mediated a rapid recovery of the dioxygenase activity in a reaction that was dependent on reductant supply. When XylT was replaced by another [2Fe-2S] ferredoxin with similar redox properties, like spinach ferredoxin, no reactivation occurred. Hence, the reactivation was XylT-specific and likely proceeded through a reduction of the active site iron of the dioxygenase. To assess this assumption, the redox state of iron at the catalytic site was monitored by EPR spectroscopy upon reactivation. Results showed that the 4-methylcatechol-inactivated C23Omt-2 displayed a signal with a S = 5/2 ground state typical for a ferric ion. Upon reactivation by a catalytic amount of XylT, this signal disappeared, indicating that FeIII was converted to FeII at the active site. Accordingly, the reactivated enzyme treated with nitric oxide yielded a prominent S = 3/2 EPR signal characteristic of a FeII-NO complex (Hugo et al., 1998). The possibility that reactivation occurred through transfer of ferrous ions from the XylT cluster to the enzyme was ruled out by EPR measurements showing that the integrity of the XylT cluster was unaltered by the reactivation reaction. Taken together, the data provided evidence that XylT mediates C23Omt-2 reactivation through reduction of the ferric ion at the enzyme active site, as illustrated in > Fig. 2. In P. putida mt-2, a NAD(P)H reductase probably serves as source of reductant in the XylT-mediated reactivation of the dioxygenase. Such an enzyme activity was detected in crude extracts by using a two-step assay involving incubation of a sample of extract with inactive C23Omt-2, XylT and NADH prior to dioxygenase assay (Jouanneau Y. and Meyer C., unpublished work). A 43-kDa protein was partially purified, but the instability of this protein precluded further characterization. Nevertheless, the XylT reductase activity was found to be 10-fold lower in a strain lacking the TOL plasmid compared to the wild-type strain mt-2, suggesting that the gene encoding the reductase was plasmid-borne. Analysis of the TOL plasmid sequence indicated that two genes might potentially encode the protein of interest. The two genes were overexpressed in E. coli, but only one of the two gene products proved capable of C23Omt-2 reactivation in the coupled assay. This protein, encoded by xylZ, was found to be the reductase component of toluate dioxygenase, a two component enzyme that catalyzes the dioxygenation of substituted benzoates (Ge et al., 2002). The xylZ gene is located in the same operon as xylT and xylE indicating that the three genes are co-transcribed.

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. Figure 2 Ferredoxin-mediated reactivation of catechol 2,3-dioxygenase. C23Omt-2 undergoes inactivation upon cleavage of 4-methylcatechol. XylT reductively reactivates C23Omt-2, using electrons provided by XylZ, which also serves as reductase for toluate dioxygenase.

Our data provide evidence that the XylZ protein has a dual function, since it acts as a reductase associated with toluene dioxygenase on the one hand, and as a XylT reductase in the specific reactivation of C23Omt-2, on the other hand (> Fig. 2).

4

Ferredoxin-Mediated Reactivation of Catechol Dioxygenases in Bacteria Degrading Other Aromatic Hydrocarbons

XylT analogues were found in Pseudomonas strains degrading naphthalene and cresols (Hugo et al., 2000) as well as in Comamonas sp. and Acinetobacter sp. degrading nitrobenzene and aniline, respectively (Tropel et al., 2002). The considered proteins had between 29 and 65% sequence identity with XylT and were purified as [2Fe-2S] ferredoxins showing similar biochemical and redox properties. Interestingly, all six proteins tested promoted the reactivation of C23Omt-2 almost as efficiently as XylT. Moreover, the analogues could functionally replace XylT in vivo, as demonstrated by complementation experiments where the introduction of the xylT-like gene in trans in a xylT mutant restored the ability of transconjugants to grow on p-methylbenzoate (Hugo et al., 2000). Like XylT, the analogues were basic proteins, suggesting that positive charges at the surface of the proteins might be involved in the interaction with catechol dioxygenase. The interactions between XylT, its analogues and C23Omt-2 were investigated by cross-linking experiments using 1-ethyl-3(3-dimethylaminopropyl)carbodiimide (EDC). A 1:1 complex was formed between one subunit of the dioxygenase and one molecule of ferredoxin, even for heterologous pairs of proteins. The formation of covalent complex was

Oxidative Inactivation of Ring-Cleavage Extradiol Dioxygenases

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affected by ionic strength, indicating that electrostatic forces were involved in the ferredoxindioxygenase interactions (Hugo et al., 2000; Tropel et al., 2002). Further studies suggested that the XylT-C23Omt-2 complex resulted from the covalent bonding between a primary amine of at least one XylT residue and a carboxylate from one acidic residue of the dioxygenase. Since there are only three lysines in the XylT sequence, replacement of each of the lysyl residues by either a glutamic acid or a glutamine was undertaken by side-directed mutagenesis (Jouanneau Y. and Meyer C., unpublished work). Results showed that variants bearing a single mutation at any of the three lysine positions formed a cross-linked complex with the dioxygenase, whereas two proteins bearing a double mutation (K34Q/K71E and K34E/ K71E) failed to form a covalent complex with C23Omt-2. This indicated that either one of the K34 or K71 residues of XylT may interact with a corresponding acidic residue of the dioxygenase. Nevertheless, all the constructed variants were found to be competent in C23Omt-2 reactivation in vitro, suggesting that electrostatic forces may not be essential for productive interaction between the two partner proteins. P. putida GJ31, a bacterium able to grow on chlorobenzene, contains a chlorocatechol dioxygenase which catalyzes the meta-cleavage of 3-chlorocatechol, in contrast to most other extradiol dioxygenases, which are inactivated by this substituted catechol (Kaschabek et al., 1998). The gene encoding this dioxygenase (cbzE) is preceded by a xylT-like gene called cbzT. The same is true for other chlorobenzene-degrading strains using the meta-cleavage pathway (Go¨bel et al., 2004). The CbzE enzyme was found to undergo inactivation during catalysis of 4-methylcatechol cleavage, suggesting that the loss of activity resulted from an oxidation of the active site iron as shown for C23Omt-2. The CbzT gene product was purified as a [2Fe-2S] ferredoxin, which efficiently reactivated CbzE in vitro, most likely by restoring the ferrous redox state at the enzyme catalytic site (Tropel et al., 2002). Curiously, while three ferredoxin analogues promoted CbzE reactivation, XylT was inefficient and failed to form a covalent complex with CbzE upon incubation with EDC. On the other hand, CbzT was competent in the reactivation of C23Omt-2 and could be cross-linked to that enzyme (Tropel et al., 2002).

5

Occurrence of xylT-Like Genes in Bacterial Genomes

A BLAST search for xylT analogues in bacterial genomes currently available in the NCBI databases yielded 53 sequences with scores above 32. All sequences share a common 4 cysteine motif for cluster ligation and exhibit a basic character (calculated pI > 7.5). Most of the retrieved genes were found in the genomes of members of the alpha, beta and gamma subgroups of Proteobacteria, and often lie close to a gene encoding a catechol dioxygenase. In sphingomonads, however, the xylT-like gene is located several kb away from a xylE gene, and no xylZ gene was detected in the large cluster of catabolic genes involved in aromatic and polyaromatic hydrocarbon biodegradation (Pinyakong et al., 2003). Surprisingly, xylT-like genes are also found in bacteria not known for their ability to degrade aromatic compounds, like the nitrogen fixing species Azotobacter vinelandii (GeneBank acc. number EAM07766) and Azoarcus sp. BH72 (GeneBank acc. number CAL95062), the latter strain being a grass endophyte. These strains contain elsewhere in their genomes at least two copies of catechol dioxygenase genes closely related to those involved in alkylphenol degradation. xylT-like genes are apparently absent from the bacterial genomes of species belonging to Actinomycetes or Bacillales, numerous members of which are endowed with the ability to degrade aromatic

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hydrocarbons. For example, Rhodococcus RHA1 has a versatile metabolism and can degrade various aromatic hydrocarbons including polychlorobiphenyls thanks to wide range of catabolic enzymes, including catechol dioxygenases (McLeod et al., 2006). Since such bacteria do not have XylT analogues, they must have evolved alternate strategies to cope with the intrinsic instability of ring-cleavage extradiol dioxygenase. In this respect, it has been observed that DHBD from B. xenovorans LB400, once inactivated by 3-chlorocatechol in vivo, recovered full activity upon removal of the inhibitor. Reactivation took about half an hour, occurred in the presence of an inhibitor of protein synthesis and was independent of a XylT-like ferredoxin (Vaillancourt et al., 2002).

6

Research Needs

The efficient and specific ferredoxin-dependent repair system described above seems to be restricted to a limited number of extradiol dioxygenases, essentially in Proteobacteria. This system is beneficial to host cells in that it extends the range of aromatic substrates they can exploit (Polissi and Harayama, 1993). Therefore, genetic transfer of xylTor one of its analogues into bacterial degraders other than Proteobacteria might be a valuable a strategy to improve the degradation efficiency of aromatic pollutants. Alternatively, search for bacterial strains producing extradiol dioxygenases more resistant to oxidative inactivation may lead to degraders with improved capabilities.

Acknowledgments I thank C. Meyer for technical assistance and helpful discussions, and J. C. Willison for critical reading of the manuscript.

References Bartels I, Knackmuss HJ, Reineke W (1984) Suicide inactivation of catechol 2,3-dioxygenase from Pseudomonas putida mt-2 by 3-halocatechols. Appl Environ Microbiol 47: 500–505. Cerdan P, Wasserfallen A, Rekik M, Timmis KN, Harayama S (1994) Substrate specificity of catechol 2,3-dioxygenase encoded by TOL plasmid pWW0 of Pseudomonas putida and its relationship to cell growth. J Bacteriol 176: 6074–6081. Dai S, Vaillancourt FH, Maaroufi H, Drouin N, Neau DB, Snieckus V, Bolin JT, Eltis LD (2002) Identification and analysis of a bottleneck in PCB biodegradation. Nat Struct Biol 9: 934–939. Ge Y, Vaillancourt FH, Agar NY, Eltis LD (2002) Reactivity of toluate dioxygenase with substituted benzoates and dioxygen. J Bacteriol 184: 4096–4103. Go¨bel M, Kranz OH, Kaschabek SR, Schmidt E, Pieper DH, Reineke W (2004) Microorganisms degrading chlorobenzene via a meta-cleavage pathway

harbor highly similar chlorocatechol 2,3-dioxygenaseencoding gene clusters. Arch Microbiol 182: 147–156. Han S, Eltis LD, Timmis KN, Muchmore SW, Bolin JT (1995) Crystal structure of the biphenyl-cleaving extradiol dioxygenase from a PCB-degrading pseudomonad. Science 270: 976–980. Harayama S, Kok M, Neidle EL (1992) Functional and evolutionary relationships among diverse oxygenases. Annu Rev Microbiol 46: 565–601. Harayama S, Rekik M (1990) The meta cleavage operon of TOL degradative plasmid pWW0 comprises 13 genes. Mol Gen Genet 221: 113–120. Hugo N, Armengaud J, Gaillard J, Timmis KN, Jouanneau Y (1998) A novel [2Fe-2S] ferredoxin from Pseudomonas putida mt-2 promotes the reductive reactivation of catechol 2,3-dioxygenase. J Biol Chem 273: 9622–9629. Hugo N, Meyer C, Armengaud J, Gaillard J, Timmis KN, Jouanneau Y (2000) Characterization of three

Oxidative Inactivation of Ring-Cleavage Extradiol Dioxygenases XylT-like [2Fe-2S] ferredoxins associated with catabolism of cresols or naphthalene: Evidence for their involvement in catechol dioxygenase reactivation. J Bacteriol 182: 5580–5585. Kaschabek SR, Kasberg T, Muller D, Mars AE, Janssen DB, Reineke W (1998) Degradation of chloroaromatics: purification and characterization of a novel type of chlorocatechol 2,3-dioxygenase of Pseudomonas putida GJ31. J Bacteriol 180: 296–302. Kita A, Kita S, Fujisawa I, Inaka K, Ishida T, Horiike K, Nozaki M, Miki K (1999) An archetypical extradiolcleaving catecholic dioxygenase: the crystal structure of catechol 2,3-dioxygenase (metapyrocatechase) from Pseudomonas putida mt-2. Structure 7: 25–34. Kovaleva EG, Lipscomb JD (2007) Crystal structures of Fe2+ dioxygenase superoxo, alkylperoxo, and bound product intermediates. Science 316: 453–457. Mars AE, Kasberg T, Kaschabek SR, van Agteren MH, Janssen DB, Reineke W (1997) Microbial degradation of chloroaromatics: use of the meta-cleavage pathway for mineralization of chlorobenzene. J Bacteriol 179: 4530–4537. McLeod MP, Warren RL, Hsiao WWL, Araki N, Myhre M, Fernandes C, Miyazawa D, Wong W, Lillquist AL, Wang D, Dosanjh M, Hara H, Petrescu A, Morin RD, Yang G, Stott JM, Schein JE, Shin H, Smailus D, Siddiqui AS, Marra MA, Jones SJM, Holt R, Brinkman FSL, Miyauchi K, Fukuda M, Davies JE, Mohn WW, Eltis LD (2006) The complete genome of Rhodococcus sp. RHA1 provides insights into a catabolic

19

powerhouse. Proc Natl Acad Sci USA 103: 15582–15587. Pinyakong O, Habe H, Omori T (2003) The unique aromatic catabolic genes in sphingomonads degrading polycyclic aromatic hydrocarbons (PAHs). J Gen Appl Microbiol 49: 1–19. Polissi A, Harayama S (1993) In vivo reactivation of catechol 2,3-dioxygenase mediated by a chloroplast-type ferredoxin: a bacterial strategy to expand the substrate specificity of aromatic degradative pathways. EMBO J 12: 3339–3347. Riegert U, Burger S, Stolz A (2001) Altering catalytic properties of 3-chlorocatechol-oxidizing extradiol dioxygenase from Sphingomonas xenophaga BN6 by random mutagenesis. J Bacteriol 183: 2322–2330. Shu L, Chiou YM, Orville AM, Miller MA, Lipscomb JD, Que L Jr. (1995) X-ray absorption spectroscopic studies of the Fe(II) active site of catechol 2,3dioxygenase. Implications for the extradiol cleavage mechanism. Biochemistry 34: 6649–6659. Tropel D, Meyer C, Jouanneau Y (2002) Ferredoxinmediated reactivation of the chlorocatechol 2,3dioxygenase from Pseudomonas putida GJ31. Arch Microbiol 177: 345–351. Vaillancourt FH, Bolin JT, Eltis LD (2006) The ins and outs of ring-cleaving dioxygenases. Crit Rev Biochem Mol Biol 41: 241–267. Vaillancourt FH, Labbe G, Drouin NM, Fortin PD, Eltis LD (2002) The mechanism-based inactivation of 2,3-dihydroxybiphenyl 1,2-dioxygenase by catecholic substrates. J Biol Chem 277: 2019–2027.

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20 Structure–Function Relationships and Engineering of Haloalkane Dehalogenases J. Damborsky* . R. Chaloupkova . M. Pavlova . E. Chovancova . J. Brezovsky Loschmidt Laboratories, Institute of Experimental Biology and National Centre for Biomolecular Research, Masaryk University, Brno, Czech Republic *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1082

2 Structure of HLDs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1083 2.1 Catalytic Residues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1083 2.2 Active Site and Tunnels . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1084 3 Function of HLDs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1085 3.1 Catalytic Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1085 3.2 Substrate Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1086 4 4.1 4.2 4.3 4.4 4.5

Engineering of HLDs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1086 Mutants with Modified Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1086 Mutants with Modified Thermostability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1090 Mutants with Modified Substrate Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1090 Mutants with Modified Enantioselectivity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1092 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1093

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_76, # Springer-Verlag Berlin Heidelberg, 2010

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20

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

Abstract: The structure–function relationships for haloalkane dehalogenases, representing one of the best characterized families of enzymes involved in degradation of halogenated compounds is described. A substantial amount of mechanistic and structural information is currently available on haloalkane dehalogenases, providing good theoretical framework for their modification by protein engineering. Examples of constructed mutants include variants with modified: (1) activity, (2) thermostability, (3) substrate specificity and (4) enantioselectivity. Some variants carried mutations in the tunnels, connecting the buried active site with surrounding solvent, rather in the active site itself. Mutagenesis in the residues lining the protein tunnels represents a new paradigm in protein engineering.

1

Introduction

Haloalkane dehalogenases (HLDs, EC 3.8.1.5) are bacterial enzymes cleaving a carbon–halogen bond in halogenated hydrocarbons. The very first HLD was isolated from Xanthobacter autotrophicus GJ10 in 1985 (Keuning et al., 1985) and served as a paradigm for carbon– halogen bond cleavage in halogenated aliphatic hydrocarbons. Since then, a number of newly isolated and biochemically characterized HLDs grown to 14 enzymes. HLDs have been isolated from bacteria colonizing contaminated environments (Janssen et al., 1988; Keuning et al., 1985; Kumari et al., 2002; Nagata et al., 1997; Poelarends et al., 1998; Poelarends et al., 1999; Sallis et al., 1990; Scholtz et al., 1987; Yokota et al., 1987), but interestingly also from pathogenic organisms (Jesenska et al., 2000; Jesenska et al., 2002; Jesenska et al., 2005). Phylogenetic analysis revealed that the HLD family can be divided into three subfamilies denoted HLD-I, HLD-II and HLD-III, of which HLD-I and HLD-III are predicted to be sister groups (Chovancova et al., 2007). A substantial amount of mechanistic and structural information is currently available on HLDs. The unique tertiary structures were determined by protein crystallography for DhlA, isolated from X. autotrophicus GJ10 (Franken et al., 1991), DhaA from Rhodococcus sp. TDTM0003 (Newman et al., 1999), LinB from Sphingobium japonicum UT26 (Marek et al., 2000), DmbA from Mycobacterium tuberculosis H37Rv (Mazumdar et al., 2008) and DbjA from Bradyrhizobium japonicum USDA110 (Prokop et al., 2009), whereas more then thirty crystal structures of protein-ligand complexes of HLDs are available in the Protein Data Bank (Supplementary Table S1). The structure and reaction mechanism of HLDs (> Fig. 1) has been studied in detail by using protein crystallography (Liu et al., 2007; Marek et al., 2000; Mazumdar et al., 2008; Newman et al., 1999; Oakley et al.,

. Figure 1 General scheme of the reaction mechanism of HLDs. Alkyl-enzyme intermediate is formed in the first reaction step by nucleophilic attack of carboxylate oxygen of an aspartate group on the carbon atom of the substrate. This intermediate is in the second reaction step hydrolyzed by an activated water molecule, yielding a halide ion, a proton, and an alcohol as the products. Enz – enzyme.

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

20

2002; Oakley et al., 2004; Ridder et al., 1999; Streltsov et al., 2003; Verschueren et al., 1993a,b,c), site-directed mutagenesis (Pries et al., 1995a,b; Bohac et al., 2002; Chaloupkova et al., 2003; Hynkova et al., 1999; Krooshof et al., 1997; Pavlova et al., 2007; Schanstra et al., 1997; Schindler et al., 1999), enzyme kinetics (Bosma et al., 2003; Prokop et al., 2003; Schanstra and Janssen, 1996; Schanstra et al., 1996a,b) and molecular modeling (Banas et al., 2006; Bohac et al., 2002; Damborsky et al., 1997a,b; Damborsky et al., 1998; Damborsky et al., 2003; Devi-Kesavan and Gao, 2003; Hur et al., 2003; Kahn and Bruice, 2003; Kmunicek et al., 2001; Kmunicek et al., 2003; Kmunicek et al., 2005; Lau et al., 2000; Lightstone et al., 1998; Maulitz et al., 1997; Nam et al., 2004; Negri et al., 2007; Olsson and Warshel, 2004; Otyepka and Damborsky, 2002; Otyepka et al., 2008; Shurki et al., 2002; Silberstein et al., 2003; Soriano et al., 2003; Soriano et al., 2005). The number of practical applications employing HLDs are increasing with growing knowledge of their properties and structure–function relationships. HLDs can find their use in the bioremediation of environmental pollutants (Stucki and Thuer, 1995), biosensing of toxic chemicals (Campbell et al., 2006), industrial biocatalysis (Janssen, 2007; Prokop et al., 2004; Swanson, 1999), decontamination of warfare agents (Prokop et al., 2005; Prokop et al., 2006), as well as cell imaging and protein analysis (Los and Wood, 2007).

2

Structure of HLDs

HLDs structurally belong to the a/b-hydrolase superfamily (Nardini and Dijkstra, 1999; Ollis et al., 1992). The proteins in this superfamily do not possess obvious sequence similarity, even though they have diverged from a common ancestor. The three-dimensional structure of HLDs is composed of two domains: (i) the a/b-hydrolase main domain, strictly conserved in various members of the a/b-hydrolase superfamily and (ii) the helical cap domain, variable in terms of number and the arrangement of secondary elements (> Fig. 2). The a/b-hydrolase fold is made mostly up of an eight-stranded parallel b-sheet which is flanked by a-helices and serves as a scaffold for the catalytic residues (Verschueren et al., 1993c). The cap domain is composed of several helices connected by loops. The cap domain is inserted to the main domain after the b-strand 6 and determines the substrate specificity (Kmunicek et al., 2001; Pries et al., 1994).

2.1

Catalytic Residues

The catalytic residues of HLDs always constitute a catalytic pentad: a nucleophile, a base, a catalytic acid (together a catalytic triad), and a pair of halide-stabilizing residues (> Fig. 2). The composition of the catalytic pentad is not conserved among different subfamilies: AspHis-Asp + Trp-Trp in subfamily HLD-I, Asp-His-Glu + Asn-Trp in subfamily HLD-II and Asp-His-Asp + Asn-Trp in subfamily HLD-III (Chovancova et al., 2007). The nucleophile is always located on a very sharp turn, known as the nucleophile elbow, where it can be easily approached by the substrate and the catalytic water molecule. The geometry of the nucleophile elbow also contributes to the formation of the oxyanion-binding site, which is needed to stabilize the negatively charged transition state that occurs during hydrolysis (Verschueren et al., 1993c). This oxyanion hole is formed by two backbone nitrogen atoms: the first is from the residue directly next to the nucleophile, while the second is located between strand b3 and

1083

1084

20

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

. Figure 2 Molecular topology (a) and tertiary structure (b) of HLDs. a/b-hydrolase fold domain (white) and the specificity-determining cap domain (black) are distinguished. A nucleophile, a base and the first halide-stabilizing residue are conserved (filled symbols), whereas the catalytic acid and the second halide-stabilizing residue are variable among HLDs (empty symbols).

helix a1 (> Fig. 2). Catalysis proceeds by the nucleophilic attack of the carboxylate oxygen of an aspartate group on the carbon atom of the substrate, yielding displacement of the halogen as a halide, and by formation of a covalent alkyl-enzyme intermediate (> Fig. 1). The alkylenzyme intermediate is subsequently hydrolyzed by a water molecule that is activated by a histidine. A catalytic acid stabilizes the charge developed on the imidazole ring of the histidine during the hydrolytic half reaction.

2.2

Active Site and Tunnels

The active site of HLDs is either a hydrophobic cavity (in DhlA) or hydrophobic pocket (in DhaA, LinB, DmbA, and DbjA) located at the interface of the main domain and the cap domain. The only polar groups localized in the active sites of HLDs are the residues of the catalytic triad. The active sites of HLDs differ in their size and accessibility to the solvent (> Fig. 3). The active site pockets can have as much as four times difference in volume: DhlA < ˙ DhlA < LinB < DmbA < DbjA. The active site cavity of DhlA is deeply buried in the protein ˙ core with limited accessibility to water molecules through a very narrow tunnel (Verschueren et al., 1993a), the active site pockets of DhaA and LinB are more accessible via the main tunnel and the slot tunnel (Petrek et al., 2006), while the pockets of DmbA and DbjA are the most exposed to solvent via the wide main tunnel and the slot tunnel (Prokop et al., 2008). These tunnels connect a hydrophobic active site with surrounding solvent and represent a very important structural feature of HLDs (Marek et al., 2000). The size, shape, physico-chemical properties, and dynamics of the tunnels are one of the determinants of activity and substrate specificity in HLDs. Tunnels play an important role during the following steps of the catalytic cycle: (1) binding of a substrate, (2) binding of catalytic water, (3) release of a halide ion, and (4) the release of an alcohol. The tunnels in HLDs can be either permanent or ligand-induced

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

20

. Figure 3 Anatomy of the active sites and tunnels in HLDs. The position of buried active site and two tunnels within protein structure is schematized in (a), where (1) denotes an active site, (2) denotes a main tunnel, and (3) denotes a slot tunnel. A surface of the active site and the tunnels is represented by wire in DhlA (b), DhaA, (c) LinB (d), DmbA (e), and DbjA (f).

(Klvana et al., 2009). The permanent tunnels are observable in the ligand-free crystal structures, while the ligand-induced tunnels are only seen in the crystal structures of the proteinligand complexes and in molecular dynamic trajectories. The solvation and desolvation of the active sites of HLDs through these tunnels is a very dynamical process due to high flexibility of the cap domain (Negri et al., 2007; Otyepka and Damborsky, 2002).

3

Function of HLDs

3.1

Catalytic Activity

A comparison of the kinetic mechanism of DhlA (Schanstra et al., 1996a), DhaA (Bosma et al., 2003) and LinB (Prokop et al., 2003) determined by transient kinetics reveals overall similarity (> Scheme 1; > Table 1). The binding of the substrate and the cleavage of the carbon–halogen bond are fast steps, resulting in the accumulation of the alkyl-enzyme intermediate for all three enzymes. The main and the very important difference in kinetic mechanism is in the ratelimiting step. The halide release is the predominant rate-limiting step for dehalogenation of 1,2-dichloroethane and 1,2-dibromoethane by DhlA (Schanstra and Janssen, 1996), liberation of alcohol for dehalogenation of 1,3-dibromopropane by DhaA (Bosma et al., 2003) and hydrolysis of the alkyl-enzyme intermediate for dehalogenation of 1-chlorohexane and bromocyclohexane by LinB (Prokop et al., 2003). The observation of different rate-limiting

1085

1086

20

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

. Scheme 1 Kinetic mechanism of HLDs. E – enzyme, RX – substrate (halogenated alkane), E.RX – enzyme-substrate complex, E-R.X – alkyl-enzyme intermediate, E.X-.ROH – enzyme-product complex, X - halide product, ROH – alcohol product. kx – kinetic constant of an individual catalytic step.

steps for three enzymes from the same protein family demonstrates that extrapolation of this important catalytic property from one enzyme to another can be misleading even for evolutionary closely related proteins.

3.2

Substrate Specificity

The HLDs are broad specificity enzymes. The set of substrates converted by HLDs consists of over a hundred chemical individuals – chlorinated, brominated and iodinated compounds; haloalkanes, haloalkenes, haloalcohols, halohydrins, haloethers, haloesters, haloacetamides, haloacetonitriles, and cyclohaloalkanes (Damborsky et al., 2001). Statistical analysis of substrate specificity profiles revealed the presence of several different specificity groups within this protein family (Damborsky et al., 1997c). Substrate specificity of HLDs is primarily determined by the structure of the cap domain (> Table 2) and can be predicted from the statistical models employing three-dimensional structures of enzyme-substrate complexes. These complexes can be prepared by computer modeling and quantitatively analyzed by using multivariate data analysis (Kmunicek et al., 2003; Kmunicek et al., 2005). Analysis of four family members revealed that only a very limited fraction of the residues (85% of variance in Michaelis constants Km. Van der Waals interactions with the residues of the first shell dominate substrate recognition in all studied HLDs (> Fig. 4). The residues of the tunnels contribute to substrate binding in LinB, DmbA, and DbjA, but not DhlA, due to low accessibility of the active site in DhlA (Brezovsky et al., unpublished).

4

Engineering of HLDs

4.1

Mutants with Modified Activity

1,2,3-trichloropropane (TCP) is a toxic non-natural compound released into the environment as a result of its manufacture, formulation, and use as a solvent and extractive agent. TCP has been detected in low concentrations in surface, drinking and ground water, with a half-life estimated to extend up to a hundred years under groundwater conditions (Yujing and Mellouki, 2001). TCP is very resistant to natural biodegradation under aerobic conditions. No natural strains, which are able to metabolize TCP have yet been isolated, opening the possibility for the construction of such a strain by genetic engineering. Construction of a dehalogenase enzyme with improved conversion of TCP is an essential step towards engineering a TCP-degrading strain. Bosma et al., (2002) applied DNA shuffling and error prone PCR on the dhaA gene to

1,2-dibromoethaned 250

DhlA W175Y

5555

1,2-dichloroethaneb

DhlA V226A

>200

bromocyclohexane

1-chlorohexaneg

LinB wt

LinB wt

b

240  44 117  5

>450

g

–

–

14.8  0.7 3.9 ± 0.6i

>75h

9.5  1

0.4  1 3.2 ± 0.2

–

1.1  0.4 2.5 ± 0.07 –

–

>40

>500

chlorocyclohexaneg

LinB wt

–

300  60

60–300

1,3dibromopropanef

DhaA wt

–

4.5 ± 1

1,2-dichloroethane

DhlA F172W

50  10h

8 ± 2h

9±2

–

14  1

>10

0.8  0.1 14  3

–

10000

c

2222

1,2-dichloroethanea

700

1,2-dibromoethane

h

16  2h

75  25h

h

h

43  10

4 ± 1.5

(s )

k4 1

8 ± 0.7

9 ± 1.5

12 ± 3

10  2

50  10

30  5

k3 (s )

1

0.55 ± 0.05 –

60  20

DhlA D260N + N148E

e

– –

>130

>27 110

–

(s )

(mM)

DhlA wt



1,2-dibromoethanec 63

DhlA F172W

Determined at pH 8.2 and 30 C (Schanstra et al., 1996a) Determined at pH 8.2 and 30 C (Schanstra et al., 1997) c Determined at pH 8.2 and 30 C (Schanstra et al., 1996b) d Determined at pH 8.2 and 30 C (Krooshof et al., 1998) e Determined at pH 8.2 and 30 C (Krooshof et al., 1997) f Determined at pH 9.4 and 30 C (Bosma et al., 2003) g Determined at pH 8.6 and 37 C (Prokop et al., 2003) h Halide release i Alcohol release

a

–

70  15

1,2-dibromoethane

DhlA V226A

b

k-2 1

(s )

1,2-dibromoethane

k2

DhlA wt

a

1

Substrate

Enzyme

KS

. Table 1 Kinetic constants of DhlA, DhaA and LinB and their mutants. Rate-limiting steps are in bold

16

23

221

5

5130

1500

530

430

60

25

33

10

(mM)

Km

2.6

1.8

0.1

3.7

2.9

3.8

3.3

0.35

5.8

5.9

8.2

3

(s )

1

kcat

0.16

0.08

0.0005

0.54

0.0006

0.0025

0.0062

0.0008

0.0008

0.24

0.25

0.3

(mM 1.s 1)

kcat/Km

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

20 1087

1088

20

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

. Table 2 Structure-specificity relationships of HLDs with known tertiary structure

Enzyme

Cap domain

Active sitea

Native substrate

Preferred substrates

DhlA

1

small b terminally halogenated

DhaA

2.5

largec terminally halogenated vicinally halogenated b-halogenated

LinB

3

largec terminally halogenated vicinally halogenated b-halogenated cyclic

DmbA

3.5

Unknown

largec monosubstituted terminally halogenated b-halogenated cyclic

DbjA

4

Unknown

largec terminally halogenated vicinally halogenated b-halogenated b-methylated cyclic

a

Relative volume Length up to C3 c Length at least C6 b

improve the kinetic properties of DhaA for TCP conversion. The evolved dehalogenase mutant, C176Y + Y273F, was 3.5-times more active towards TCP than the wild type enzyme. Another variant of DhaA (Gray et al., 2001) also carried a substitution at position 176. This random variant of DhaA, G3D + C176F, was obtained by in vitro evolution and showed a 4-fold improvement in activity with TCP, relative to the wild type enzyme. Pavlova et al., (2009) combined advanced computer modeling with directed evolution and obtained twenty five unique protein variants with higher activities towards TCP than the wild type enzyme. The best mutant carried five single-point mutations and demonstrated 32-times higher

. Figure 4 Interactions important for recognition of substrates by the active sites of HLDs: DhlA (a), LinB (b), DmbA (c), and DbjA (d). The interactions are labeled according to chemical character (van der Waals, v; electrostatics, e), ordered by their importance for the multivariate model and colored according to their localization in the first shell (blue), second shell (red) and tunnel (yellow).

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

20 1089

1090

20

Structure–Function Relationships and Engineering of Haloalkane Dehalogenases

activity (26-times higher catalytic efficiency) when compared to the natural wild type enzyme. The ‘‘hot spot’’ residues for saturated mutagenesis were selected by Random Acceleration Molecular Dynamics (Luedemann et al., 2000), simulating the release of the product from the enzyme active site. Interestingly, mutagenesis targeted the access tunnels rather than the active site. These tunnels connect the buried active site cavity with the surrounding solvent and the enhanced rate with TCP appears to be due to the absence of water molecules in the active site cavity promoting formation of an activated complex (> Fig. 5). Efficiently catalyzed reaction steps are followed by solvation of the active site by water molecules. Waters are attracted to the cavity from bulk solvent due to the presence of charged ion and assist release of products.

4.2

Mutants with Modified Thermostability

The HLDs represent a class of enzymes with a high potential for biocatalysis (Janssen, 2007). Performance of the biocatalytic process is a combination of the reaction rate of a biocatalyst and its stability. According to the Arrhenius relationships, the rate of the enzymatic reaction will approximately double for every 10 C increase in temperature. Gray et al., (2001) attempted to improve stability of DhaA at higher temperatures to develop efficient biocatalytic process for the conversion of halogenated alkanes to halohydrin products (Swanson, 1999). They used a directed evolution technique called the Gene Site Saturation Mutagenesis (Kretz et al., 2004), which theoretically allowed all single site mutants to be sampled, in combination with high-throughput screening methods. Thermostability of parental dehalogenases and evolved mutants was measured by assaying activity at elevated temperatures. Eight single point mutations were discovered to be scattered along the protein sequence. This had considerable effects on enzymes thermostability (> Fig. 6). A combination of all of these mutations yielded a variant D78G + F80S + T148L + G171Q + I209L + N227T + W240Y + P291A with a 30,000-times longer half-life at 55 C, and an increase in Tm to 8 C. Stabilization of an a-helix by the mutation T148L, which is responsible for the formation of an additional H-bond between the serine hydroxy group and an acceptor in F80S mutant, were important factors for the enhanced stability of enzymes. Effects of other mutations, including I209L and P291A, with the largest contribution towards improved thermostability were more difficult to explain. Three mutations N227T, W240Y, and P291A did not affect melting temperature, although did contribute to the increase of half life. A plausible mechanistic explanation is that these three mutations increase the possibility that the protein will refold more efficiently after denaturation. The complexity of the results demonstrate that our understanding of structural basis of protein stabilization is still limited as it would be very difficult, if not impossible, to design these stabilizing mutations rationally.

4.3

Mutants with Modified Substrate Specificity

Investigation of the evolution of enzymes by selecting spontaneous mutants that convert a xenobiotic compound, represents a unique opportunity to observe how new substrate specificities evolve in Nature. 1,2-Dichloroethane (DCE) is a non-natural compound whose production and emission to the biosphere started in 1922. It is unlikely that sufficient selective pressure to evolve a complementary enzyme existed before this date (Pries et al., 1994) and the enzymes participating in the degradation of DCE must have undertaken recent evolutionary

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. Figure 5 Water accessibility in the wild type DhaA (a) and its five-fold mutant I135F + C176Y + V245F + L246I + Y273F (b) with enhanced activity towards TCP. Spheres show the positions of water molecules inside the protein for snapshots collected during the molecular dynamics simulation. ˚ and are centered on the oxygen atoms of the water molecules. The spheres have a radius of 0.5 A The mutants were designed by Random Expulsion Molecular Dynamics simulations and constructed by site-saturated mutagenesis (Pavlova et al., 2009).

. Figure 6 Substitutions accumulated in the eight-point mutant of DhaA with enhanced thermostability (D78G + F80S + T148L + G171Q + I209L + N227T + W240Y + P291A). All but one substitution (I209L) are located on the protein surface. The mutants were obtained by Gene Site Saturated Mutagenesis (Gray et al., 2001).

adaptation. The first step in the utilization of DCE by soil bacterium X. autotrophicus GJ10 is catalyzed by DhlA, hydrolyzing this short-chain (C2) chloroalkane to the corresponding alcohol, which can further serve as source of carbon and energy for growth. In a fascinating laboratory evolution experiment, Pries et al., (1994) expressed DhlA in a strain of Pseudomonas that grows on long-chain (C6) alcohols and selected 12 independent mutants that utilize

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. Figure 7 Spontaneous substitutions (D170H and P168S), deletion (D164–174) and insertions (∇172–174, ∇152–153 and ∇145–154) localized in the cap domain of DhlA mutants with relaxed substrate specificity. The residues making the salt bridge D170-K261 between the main domain and the cap domain are shown in ball and stick. The mutants were obtained from an in vivo adaptation experiment (Pries et al., 1994).

1-chlorohexane. These mutants were obtained after 4 weeks in batch cultivations that contained a mixture of 1-chlorobutane and 1-chlorohexane as the sole carbon sources. Sequencing of evolved genes revealed six mutant dehalogenases with relaxed substrate specificities: ∇145–154, ∇152–153, D164–174, P168S, D170H, and ∇172–174. Interestingly, none of the mutations directly affected the active site residues, with the exception of D164–174, in which the active site cavity forming residues F164 and F172 were missing. All observed mutations are located in a segment of the dhlA gene which encodes the N-terminal part of the cap domain (> Fig. 7). The mutants D164–174, P168S, D170H, and ∇172–174 carry changes that affect the structurally important salt bridge D170-K261. This salt bridge is positioned between two domains and its disruption will make the cap domain more floppy (Otyepka and Damborsky, 2002). The structural basis of the relaxed specificity in the other two mutants, ∇145–155 and ∇152–153, is more difficult to explain. The active site cavity could be enlarged due to insertions, but this is only speculation, as the residues surrounding the insertion are not in direct contact with the substrate of the wild type enzyme (Pries et al., 1994). These results present experimental evidence that the cap domain determines substrate specificity and that generation of the repeats is an important mutational event during its evolution. This evolutionary paradigm has been recently implemented into a novel directed evolution method which generates randomly repeats and deletions in vitro (Pikkemaat and Janssen, 2002).

4.4

Mutants with Modified Enantioselectivity

In response to the general awareness of the physiological and ecological advantages of the use of single enantiomers, the manufacture of enantiomerically pure compounds has become an expanding area of the fine chemicals industry. When pharmaceuticals, agrochemicals, food additives and their synthetic intermediates are marketed as single enantiomers, high

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enantiomeric purities characterized by enantiomeric excess (e.e.) >98%, are required. Pieters et al., (2001) investigated chiral recognition of haloalkane dehalogenases DhlA and DhaA. The magnitude of chiral recognition was low; a maximum E-value of 9 could be reached after structural optimization of the substrate and the development of enantioselective dehalogenases for use in industrial biocatalysis was defined as one of the major challenges of the field (Janssen, 2004). We have assayed DhaA, LinB, and DbjA for their enantioselective conversion of brominated esters and alkane substrates into chiral alcohols (Prokop et al., 2009). All three proteins possessed high enantioselectivity (>200) with a-brominated esters. DbjA additionally showed high enantioselectivity towards structurally simple molecule 2-bromopentane (E = 145), while DhaA and LinB showed only low enantioselectivity (E = 7 and E = 16, respectively). Structural analysis revealed that DbjA contains a unique surface loop in its specificity-determining domain. Deletion of this loop has led to the mutant enzyme DbjAD with a significantly lowered enantioselectivity toward 2-bromopentane (E = 58). Enantioselectivity could be re-introduced by an additional single-point mutation DbjAD + H139A (E = 120). Introduced mutations modulated anatomy and water accessibility of the main tunnel in DbjA (> Fig. 8). A hydrophobic interaction of the alkyl chain with the wall of this tunnel accompanied by desolvation seemed to be important for enantioselective discrimination of the structurally simple molecule 2-bromopentane by DbjA. Another two studied family members, DhaA and LinB, do not have this water accessible cone-like tunnel and therefore cannot efficiently discriminate enantiomers of b-brominated alkanes. These results demonstrate that enantioselectivity of an enzyme can be modulated by engineering of a protein tunnel via modification of a surface loop.

4.5

Research Needs

Isolation and biochemical characterization of new members of the HLD family continues to be of great interest. Characterization of new family members has led to new knowledge about structure–function relationships and the evolution of HLDs. These newly isolated enzymes,

. Figure 8 The deletion mutants of DbjA with modified tunnels and modulated enantioselectivity: wild type DbjA (a), D140–146 (b) and D140–146 + H139A (c). The region carrying deletion in the surface loop is shown in ribbon. The ‘‘gate-keeping’’ His/Ala139 are shown in stick. The mutants were designed based on sequence/structure comparisons and constructed by site-directed mutagenesis (Prokop et al., 2009).

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native or genetically modified, also hold great potential for practical applications which often require optimized properties: (1) high enantioselectivity with substrates that can be converted to valuable products by biocatalysis, (2) enhanced resistance to organic solvents for decontamination purposes, (3) elevated activities with specific target compounds like TCP or DCE for bioremediation, (4) broadened pH-range for biosensors, and (5) increased thermostabilities and long-term stabilities for nearly every possible application. Development of better data management and tools for analysis are needed for mechanistic studies. The amount of data on HLDs, as well as many other enzymes, is growing exponentially and these tools will assist in the extraction of knowledge from this data. For example, we have only started to understand the importance of tunnels in HLDs for (de)solvation and the exchange of ligands between the active site and the surrounding environment, even though these processes are essential for function of proteins with buried active sites. The greatest challenge in the research of HLDs is the identification of their biological role. The genes coding for HLDs are widely distributed among various bacterial species, including the tissue-colonizing organisms, e.g., Mycobacterium tuberculosis or Mycobacterium bovis. The number of genes annotated as HLDs by sequence similarity in genomic and proteomic databases is growing. Though for many proteins encoded by these genes, dehalogenating activity has not yet been confirmed experimentally and their natural function in host organisms remains unknown.

Acknowledgments Financial support of the Ministry of Education, Youth and Sports of the Czech Republic via LC06010 (J. Brezovsky, E. Chovancova, M. Pavlova) and MSM0021622412 (J. Damborsky, R. Chaloupkova) is gratefully acknowledged.

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reaction in haloalkane dehalogenase. J Am Chem Soc 127: 1946–1957. Streltsov VA, Prokop Z, Damborsky J, Nagata Y, Oakley AJ, Wilce MCJ (2003) Haloalkane dehalogenase LinB from Sphingomonas paucimbilis UT26: X-ray crystallographic studies of dehalogenation of brominated substrates. Biochemistry 42: 10104–10112. Stucki G, Thuer M (1995) Experiences of a large-scale application of 1,2-dichloroethane degrading microorganisms for groundwater treatment. Environ Sci Technol 29: 2339–2345. Swanson PE (1999) Dehalogenases applied to industrialscale biocatalysis. Curr Opin Biotechnol 10: 365–369. Verschueren KHG, Franken SM, Rozeboom HJ, Kalk KH, Dijkstra BW (1993a) Refined X-ray structures of haloalkane dehalogenase at pH 6.2 and pH 8.2 and implications for the reaction mechanism. J Mol Biol 232: 856–872.

Verschueren KHG, Kingma J, Rozeboom HJ, Kalk KH, Janssen DB, Dijkstra BW (1993b) Crystallographic and fluorescence studies of the interaction of haloalkane dehalogenase with halide ions. Studies with halide compounds reveal a halide binding site in the active site. Biochemistry 32: 9031–9037. Verschueren KHG, Seljee F, Rozeboom HJ, Kalk KH, Dijkstra BW (1993c) Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature 363: 693–698. Yokota T, Omori T, Kodama T (1987) Purification and properties of haloalkane dehalogenase from Corynebacterium sp. strain m15–3. J Bacteriol 169: 4049–4054. Yujing M, Mellouki A (2001) Rate constants for the reactions of OH with chlorinated propanes. Phys Chem Chem Phys 3: 2614–2617.

21 Lipolytic Enzymes from Bacteria S. Hausmann1,2 . K.-E. Jaeger1 1 Institut fu¨r Molekulare Enzymtechnologie, Heinrich-Heine-Universita¨t Du¨sseldorf, Forschungszentrum Ju¨lich, Ju¨lich, Germany [email protected] 2 Evocatal GmbH, Du¨sseldorf [email protected]

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1100

2

Structure Function Relationships of Lipolytic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . 1100

3

Distribution and Classification of Bacterial Carboxylesterases and Lipases . . . . . 1102

4

Physiological Functions of True Lipases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1102

5

Triacylglycerol Lipases (Family I) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1104

6

Esterases (Families II–VIII) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1110

7

Physiological Functions of Esterases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1110

8

Family II . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1113

9

Family III . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1114

10 Family IV . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1114 11 Family V . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1115 12 Family VI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1115 13 Family VII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1116 14 Family VIII . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1116 15 Research Needs and Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1117

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_77, # Springer-Verlag Berlin Heidelberg, 2010

1100

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Lipolytic Enzymes from Bacteria

Abstract: Lipolytic enzymes comprising carboxylesterases and lipases represent a highly diverse group of hydrolases. Presently, about 900 of these enzymes are identified to originate from bacteria. In a landmark publication which appeared 10 years ago (Arpigny and Jaeger, 1999), lipolytic enzymes were grouped into eight families based on amino acid sequence homology and physiological properties. Here, we present the current status of this classification framework into which we have included numerous novel bacterial lipolytic enzymes. We further describe their biochemical properties and characteristic structural features. Additionally, diverse physiological functions of selected lipolytic enzymes are discussed.

1

Introduction

Hydrolases (IUBMB classification EC 3) catalyze the cleavage of chemical bonds by the addition of water. Under physiological conditions, the excess of water renders these reactions irreversible for thermodynamic reasons. Hydrolases are classified into 13 subgroups based on the type of bond they act: subgroup EC 3.1 includes enzymes acting on ester bonds, EC 3.2 on glycosides, EC 3.3 on ether, and EC 3.4 on peptide bonds. The huge number of known hydrolases and their enormous diversity emphasize their biological importance. This chapter focuses on lipolytic enzymes which include carboxylesterases (EC 3.1.1.1) and true lipases (EC 3.1.1.3). Carboxylesterases act on small and usually water-soluble esters; the reaction kinetics can be described by the Michaelis-Menten equation (Brockerhoff and Jensen, 1974; Sarda and Desnuelle, 1958). In contrast, true lipases preferentially hydrolyze ester bonds of triglycerides with long-chain fatty acids which are often insoluble in water (Verger, 1997). The reaction kinetics of true lipases are characterized by sigmoid curves (Brockerhoff and Jensen, 1974) because these enzymes show a behavior called ‘‘interfacial activation.’’ At low substrate concentrations, enzyme activity is also low; however, as soon as the substrate concentration increases to a point where the substrate starts to form aggregates (critical micellar concentration), an interface is formed between hydrophobic substrate micelles and the hydrophilic water phase containing dissolved lipase. At this point, lipolytic activity rapidly increases. A structure-based explanation for this observation was deduced from the crystal structures of human pancreatic lipase and a lipase from the fungus Rhizomucor miehei. These tertiary structures showed a flexible a-helix which covered a tunnel leading to the active site. Upon contact with an interface, the a-helix termed the ‘‘lid’’ undergoes a conformational change and the active site becomes accessible to the substrate (Brzozowski et al., 1991; van Tilbeurgh et al., 1993). At the same time, the hydrophobic surface area surrounding the lipase catalytic site is increased thereby promoting the interaction of enzyme and substrate (Brzozowski et al., 1991). Later, several lipases were identified and biochemically characterized which did not show interfacial activation, e.g., the lipase LipA of B. subtilis, which does not even possess a lid, and P. aeruginosa lipase LipA which does not show interfacial activation either, although this enzyme contains a lid (Jaeger et al., 1993; Lesuisse et al., 1993; van Pouderoyen et al., 2001). Hence, the presence of a lid is presently not regarded as a structural feature distinctive between carboxylesterases and true lipases (Verger, 1997).

2

Structure Function Relationships of Lipolytic Enzymes

Lipases were classified into eight subfamilies based on amino acid sequence homology and physiological functions (Arpigny and Jaeger, 1999; Jaeger and Eggert, 2002). Also, crystal

Lipolytic Enzymes from Bacteria

21

structures of lipases from 25 different organisms were solved, among them of lipases from all bacterial subfamilies except families I.7 and I.8. (Lang et al., 1996; Meier et al., 2007; Nardini et al., 2000; Noble et al., 1993; Tyndall et al., 2002; van Pouderoyen et al., 2001). Most enzymes belong to the family of a/b-hydrolases that comprises not only esterases but a wide variety of other hydrolases like proteases, amidases, and many others (Nardini and Dijkstra, 1999). The canonical a/b hydrolase fold consists of a central, mostly parallel b-sheet of eight strands with the second strand antiparallel. The parallel strands b3–b8 are connected by a helices, which pack on either side of the central b-sheet (Nardini and Dijkstra, 1999; Ollis et al., 1992). Many a/b-hydrolases including lipolytic enzymes are also referred to as serine hydrolases (Blow, 1990) because they possess a catalytic triad consisting of Ser, His, and Asp (or Glu) residues (Brady et al., 1990; Brenner, 1988; Schrag et al., 1991). In the primary structure, these residues follow the order Ser–Asp–His. The catalytically active serine is usually located within a conserved pentapeptide with the sequence G-X-S-X-G with serine positioned at the ‘‘nucleophilic elbow,’’ a characteristic b-e-Ser-a motif that seems to be a steric prerequisite for enzymatic activity (Ollis et al., 1992). Hydrolysis of an ester bond is initiated by a nucleophilic attack of the serine residue on the carbonyl carbon-atom of the ester bond, forming a tetrahedral transition state (Semeriva et al., 1974), which is stabilized by hydrogen bonding to backbone amide residues of the so-called oxyanion hole (> Fig. 1). After the corresponding alcohol is released, a lipase–acyl complex is formed which is subsequently hydrolyzed releasing the free fatty acid and the enzyme (Jaeger and Reetz, 1998; Jaeger et al., 1999).

. Figure 1 (a) Active site of a lipolytic enzyme. Blue arrows indicate proton transfer mediated by the catalytic triad residues Asp, His, and Ser, and the nucleophilic attack of the catalytic Ser on the carbonyl carbon-atom of the substrate ester bond. Red dashed lines indicate the interaction of backbone amides forming the oxyanion hole with the substrate. (b) Active site of P. aeruginosa lipase LipA (PDB Code 1EX9) (Nardini et al., 2000). The catalytic triad residues Ser82, Asp229, and His251 are shown as sticks, the substrate (a triacylglycerol analog) is highlighted in ball and stick mode. The surface electrostatic charges are indicated in blue for positive and red for negative charges, respectively. The structure was visualized using USCF Chimera (Pettersen et al., 2004).

1101

1102

21

Lipolytic Enzymes from Bacteria

. Figure 2 Taxonomic distribution of lipases and esterases annotated in the BRENDA database. Enzyme entries were counted and classified according to their origin from archaea, bacteria, fungi, other eukaryotes, and origin not further specified.

3

Distribution and Classification of Bacterial Carboxylesterases and Lipases

Initially, 53 known lipases and carboxylesterases were classified into eight families based on their properties and sequences (Arpigny and Jaeger, 1999). Meanwhile, numerous lipolytic enzymes have been newly identified and characterized. An excellent nonredundant collection of known lipases is provided by the BRENDA database (Pharkya et al., 2003; Schomburg et al., 2004). Presently (January 2009), this database contains entries for 745 triacylglycerol lipases and 696 carboxylesterases. An overview of the taxonomical distribution of these enzymes is given in > Fig. 2. Obviously, the majority of both classes of annotated enzymes originates from microorganisms, namely bacteria and fungi, and 369 lipases and 410 esterases are annotated for genes originating from bacteria. The respective amino acid sequences were aligned using the ClustalW algorithm (Myers and Miller, 1988) and grouped into lipase families I–VIII as proposed in 1999 (Arpigny and Jaeger, 1999). Since then, the number of lipolytic enzymes within the proposed families has increased approx. sixfold and crystal structures have also been solved of representative members for most of the families.

4

Physiological Functions of True Lipases

Interestingly, the physiological functions of many bacterial lipases are still not fully understood. However, their expression often underlies a complex regulation implying that their primary role may exceed the mere hydrolysis of fats to supply the cells with putative carbon sources (see > Fig. 3). In many pathogenic bacteria belonging or related to the genus Pseudomonas, regulation of lipase gene expression was found to be quorum-sensing dependent. As this mechanism often regulates the expression of genes involved in virulence (Degrassi et al., 2008; Devescovi et al., 2007; Heurlier et al., 2004; Lewenza et al., 1999; Ulrich et al., 2004) several lipases may act as virulence factors as well. In plants, they not only hydrolyze lipid bodies and epicuticular

Lipolytic Enzymes from Bacteria

21

. Figure 3 Putative physiological functions of lipolytic enzymes with major functional areas including virulence, metabolism, and growth promotion. Further details are described in the text (CAMP: cationic antimicrobial peptide).

waxes, but also the ester bonds of xylan thereby overcoming protective barriers of the host (Khalameyzer et al., 1999). P. aeruginosa is an opportunistic pathogen that infects the respiratory tract of cystic fibrosis patients. Its lipase and phospholipase C can degrade dipalmitoyl-phosphatidylcholin

1103

1104

21

Lipolytic Enzymes from Bacteria

(DPPC) resulting in the formation of dipalmitoylglycerol and palmitate (Beatty et al., 2005). DPPC represents the main component of the host lung surfactant which is needed to reduce the surface tension of alveoli membranes to facilitate breathing (Schmidt et al., 2007). P. aeruginosa cells isolated from cystic fibrosis lungs contain LPS with an unusual lipid A composition containing up to 33% palmitate, a fatty acid which is not present in lipid A of wild-type P. aeruginosa (Ernst et al., 1999). It was also found that Salmonella strains containing palmitate in their lipid A displayed significantly increased resistance against cationic antimicrobial peptides (CAMPs) and were recognized less effective by the immune system of the host (Guo et al., 1998; Tanamoto and Azumi, 2000). Moreover, the presence of free fatty acids can result in several physiological effects hampering the host immune response including inhibition of lymphocyte proliferation and chemotaxis of human neutrophils (Buttke, 1984; Buttke and Cuchens, 1984; Hawley and Gordon, 1976; Jaeger et al., 1991; Nordstrom et al., 1991; Pourbohloul et al., 1985). Thus, lipolytic enzymes presumably aid bacteria to colonize the respiratory tract as well as to increase their resistance against the host immune defense system. The lipase of Propionibacterium acnes, a bacterium that resides in the psilobaceous follicles, most likely represents an important colonization factor. None of several lipidic substrates added to continuous cultures affected growth of this bacterium, and glycerol was only a poor carbon and energy source as compared to glucose. However, bacterial cells adhered to different lipid species, preferentially to free fatty acids (Burkhart and Burkhart, 2003; Gribbon et al., 1993). Hence, lipase-mediated hydrolysis of sebum glycerides provides free fatty acids to aid bacterial colonization. Lipolytic enzymes also play an important role in the degradation of lipidic compounds to provide carbon sources. As an example, P. aeruginosa YS-7 is able to colonize harsh ecological niches, like oil emulsions with very low water content (1%) (Shabtai, 1991). Microthrix parvicella, a filamentous bacterium populating activated sludge plants, is specialized in uptake and metabolism of lipids and fatty acids contained in the wastewater in high concentrations. This organism takes up and degrades these compounds under both aerobic and anaerobic conditions, whereas most microorganisms need aerobic conditions for uptake (Nielsen et al., 2002). It is known that the expression of different lipases can be stimulated by addition of hydrophobic compounds, e.g. the alkane hexadecane (Boekema et al., 2007; Breuil et al., 1978; Gilbert et al., 1991; Kanwar et al., 2002; Kok et al., 1996; Mahler et al., 2000; Martinez and Nudel, 2002; Shabtai and Daya-Mishne, 1992). Lipases are involved in alkane degradation although the underlying mechanisms are not fully understood. When 56 different strains were tested for their potential to metabolize hexadecane as the sole carbon source, none of the lipase-negative bacteria was able to grow. On the other hand, only half of the lipolytic strains investigated could grow on hexadecane demonstrating that lipase production alone is not sufficient to utilize alkanes as a carbon source (Breuil et al., 1978). Not surprisingly, alkanes are poor carbon sources for bacteria. In the presence of glucose, ethanol, or acetate, diauxic growth occurs and alkanes are utilized latest (Breuil et al., 1978; Makula et al., 1975). Only few bacteria can grow on alkanes as shown in a study where 61 bacterial and 28 yeast strains were tested: only 6.6% of the bacteria, but 78.6% of the yeast strains could use hexadecane as the sole carbon source (Margesin et al., 2003).

5

Triacylglycerol Lipases (Family I)

Family I combines all true lipases and consists of eight subfamilies (> Table 1). Lipases of subfamilies I.1 and I.2 can be distinguished by their amino acid sequences with enzymes

Lipolytic Enzymes from Bacteria

21

belonging to subfamily I.1 showing a high sequence similarity to the lipase of P. aeruginosa and their molecular masses range from about 30 to 32 kDa. Lipases of family 1.2 exhibit a significant similarity to the B. glumae lipase and have a slightly larger molecular mass because they possess two additional antiparallel b-strands at the protein surface that are not present in lipases of family I.1. Both subfamilies comprise lipases of gram-negative bacteria which are secreted by the type II secretion pathway, a two-step translocation mechanism that involves a large number of accessory proteins. The cytoplasmic membrane is crossed either via the Secor the Tat-pathway (Pugsley, 1993; Saier, 2006; von Heijne, 1990; Voulhoux et al., 2001; Watson, 1984) that both mediate ATP-dependent translocation into the periplasm (Driessen et al., 1998; Duong and Wickner, 1997; Economou, 1998; Hanada et al., 1994). Here, proteins fold into the active conformation before they are exported into the extracellular space (Braun et al., 1996; Koster et al., 2000; Ma et al., 2003; Pugsley, 1992; Tommassen et al., 1992). Several lipases also need the Dsb-system for the formation of disulfide bonds (Missiakas and Raina, 1997; Pugsley, 1992) and interaction with specific intermolecular chaperones, so-called lipasespecific foldases or ‘‘Lif proteins’’ (Aamand et al., 1994; Frenken et al., 1993a; Hobson et al., 1993; Ihara et al., 1995; Kok et al., 1995; Rosenau et al., 2004). Lif proteins were identified in a variety of different strains, including Burkholderia, Pseudomonas, Acinetobacter, and Vibrio species (Frenken et al., 1993a, b; Joergensen et al., 1991; Kok et al., 1995; Ogierman et al., 1997; Wohlfarth et al., 1992). Usually, they are encoded in a bicistronic operon together with the cognate lipase suggesting highly specific recognition of their target enzymes (El Khattabi et al., 1999; Shibata et al., 1998). The protein complex formed by the lipase and the Lif of B. glumae was crystallized and its structure solved (Pauwels et al., 2005, 2006). In this complex, the

. Table 1 Family I of lipolytic enzymes comprising true lipases Subfamily I.1

Origin

Accession No.

Pseudomonas aeruginosa

P26876

Pseudomonas mendocina

Q8RKT7

Rhodoferax ferrireducens

Q21T36

Vibrio cholerae

P15493, A6A105, A5EYU1, A1F3S2, A3H0H8, A2PI47, A3EDG4, A3GSF6, A1EMR8, A2P8X1, A6AE39, A6XWR7, A2PS04, A3EIQ3

Vibrio parahaemolyticus

A6B1H2

Vibrio harveyi

A6AS17

Aeromonas hydrophila

A0KFL9

Dehalococcoides sp. VS

A8CY80

Chromobacterium violaceum

Q7NUI4

… and 16 others

…

1105

1106

21

Lipolytic Enzymes from Bacteria

. Table 1 (Continued) Subfamily I.2

I.3

I.4

Origin

Accession No.

Burkholderia glumae

Q05489

Burkholderia cenocepacia

Q1BM22, A0AZ26, B1K3P3

Burkholderia multivorans

Q45VN4, A9AMF2

Burkholderia thailandensis

Q2T7L1

Pseudomonas KWI-56

P25275

Burkholderia cepacia

P22088, Q4JL88, Q6B4I1, Q0BAH5

Pseudomonas luteola

O68551

… and 7 others

…

Pseudomonas fluorescens PfO1

Q3KCS9

Pseudomonas fluorescens

P26504, P41773, Q76D26, Q76D27, Q76D28, Q76D29, Q76D31, Q76D32

Pseudomonas sp. 7323

Q2KTB3

Pseudomonas entomophila

Q1IBE9

Serratia proteamaculans

A8GDX0

Serratia marcescens

Q09KJ5, Q0MVP2, Q59932, Q59933

uncultured bacterium

A7J993

Psychrobacter sp. PR-Wf-1

A5WGV1

Bacillus subtilis

P37957

Bacillus amyloliquefaciens

A7Z124

Bacillus pumilus

A1E152, A8FGA4

Bacillus licheniformis

Q9K5F4

Bacillus sp. NK13

B0LW76

Bacillus megaterium Q8RJP5 Bacillus clausii

Q5WDN0

Lipolytic Enzymes from Bacteria

21

. Table 1 (Continued) Subfamily I.5

I.6

I.7

I.8

Origin

Accession No.

Bacillus thermocatenulatus

Q59260

Geobacillus zalihae

Q842J9

Bacillus sp. L2

Q5I4I3

Geobacillus sp. SF1

Q1L776

Bacillus stearothermophilus

A0MTM1, O66015, Q93A71, Q9L6D3

Geobacillus kaustophilus

Q5KYG5

… and 5 others

…

Staphylococcus hyicus

P04635

Staphylococcus simulans

Q84EK3

Staphylococcus xylosus

Q2TPV1

Staphylococcus epidermis

P0C0R3, Q9Z4M7, P0C0R4, Q5HKF8, Q5HKP6

Staphylococcus haemolyticus

Q9RGZ6

Staphylococcus warneri

Q5DWE2

Staphylococcus aureus

P10335, Q59811, Q79SZ7, Q2YVD0, Q2YZ74, Q5HCM7, Q5HJ48, A6TYA4, A6U555, A5IPI7, A5IW97, Q6GDD3, Q6GJZ6, Q6G604, Q6GCF1, P65288, Q99WQ6, Q8NUI5, Q8NYC2, P65289, Q7A7P2, Q2FUU5, Q2G155, A8YZE4, A8Z5H0, Q2FDJ1, Q2FJU4

Streptomyces cinnamoneus

O33969

Propionibacterium acnes

Q59644, Q6A6T8

Corynebacterium glutamicum

Q8NU60, Q8NU59

Pseudoalteromonas haloplanktis

Q3IF07

Hahella chejuensis

Q2SGZ8

Colwellia psychrerythraea

Q48AN1

Pseudoalteromonas tunicata

A4CF12

1107

1108

21

Lipolytic Enzymes from Bacteria

foldase consists of a mainly a-helical motif comprised of 11 a-helices with the N- and C-terminal regions forming minidomains built up by three helices each. These domains most likely play a key role in lipase binding to the foldase. The previously identified conserved motif in lipase-specific foldases RX1X2FDY(F/C)L(S/T)A (Rosenau et al., 2004) is located in the N-terminal binding domain. The high specificity of lipase–foldase interactions is impressively reflected by an exceptionally large pairwise interaction surface of 5378 A˚2 which is more than three times the area found for an average protein–protein interaction (1,600 A˚2 (Wodak and Janin, 2002), and a dissociation constant KD of only 5 nM (Pauwels et al., 2006, 2007). Finally, the correctly folded and processed proteins are recognized by the complex type II secretion machinery consisting of up to 14 different proteins (Sandkvist, 2001) which mediates the secretion through the bacterial outer membrane (Filloux et al., 1998; Koster et al., 2000; Tommassen et al., 1992). Further common characteristics of lipases from both subfamilies include two aspartic acid residues that mediate the coordination of a Ca2+ ion which is essential for the catalytical active structure. For the lipase of P. aeruginosa, analysis of the tertiary structure (Nardini et al., 2000) revealed the presence of a loop containing the catalytic histidine which is stabilized and oriented properly by the Ca2+. Additionally, most of these lipases contain two cysteines that form an intramolecular disulfide bond which stabilizes the folded enzyme, but is not necessary for its correct folding, its interaction with the foldase, or catalytic activity (Liebeton et al., 2001). Subfamily I.3 lipases have a molecular mass of 50–65 kDa and, in contrast to family I.1 and I.2 enzymes, are exported by the type I secretion pathway consisting of a threecomponent ATP-binding cassette transporter system (Arpigny and Jaeger, 1999). Instead of an N-terminal signal sequence, these lipases carry a C-terminal secretion signal mediating their one-step translocation into the extracellular space (Akatsuka et al., 1994; Amada et al., 2000; Kwon et al., 2002). Recently, the structure of the lipase LipA from Serratia marcescens was solved providing insight for the first time into the tertiary structure of a family I.3 enzyme (Meier et al., 2007). LipA consists of 613 amino acids of which the N-terminal 320 residues form the catalytically active lipase domain which revealed a modified a/b hydrolase fold different from the folds observed for other bacterial lipases. As a unique feature, a Ca2+ ion was identified which binds the lid helix of the enzyme and is essential for enzymatic activity. A comparison of the S. marcescens LipA structure with other lipase tertiary structures revealed that this enzyme is closer related to eukaryotic than to bacterial lipases. The C-terminal part of this lipase consists of a novel so-called b-roll sandwich motif comprising two separated b-roll domains tightly packed against each other. Lipase, like other passenger proteins secreted by the type I pathway, possesses characteristic glycine-rich repeats (GGXGXDX(U)X)n preceding the C-terminal secretion signal. This so-called RTX-signature (for repeats in toxins) mediates binding of Ca2+ ions (Welch, 2001) and may act as an intramolecular chaperone at higher calcium concentrations in the extracellular medium by providing a nucleus for folding. In addition, it may prevent early folding at low Ca2+ concentrations present in the cytoplasm (Bauche et al., 2006; Lilie et al., 2000; Meier et al., 2007; Rose et al., 1995). Subfamily I.4 lipases are the smallest triacylglycerol lipases currently identified, exhibiting a molecular mass of about 20 kDa (Arpigny and Jaeger, 1999). In this group, several lipases of the gram-positive genus Bacillus can be found, including B. licheniformis, B. subtilis and B. pumilus lipases which all contain the conserved pentapeptide sequence Ala-X-Ser-X-Gly with an Ala in position 1 as compared to the canonical sequence motif Gly-X-Ser-X-Gly.

Lipolytic Enzymes from Bacteria

21

Lipases of this family show maximal activity at pH 10.0–11.5 (Nthangeni et al., 2001). In contrast to lipases from families I.1 to I.3, stability and activity of lipase from B. pumilus B26 is independent from Ca2+ ions and the enzyme does not contain any cysteine residues (Kim et al., 2002; Nthangeni et al., 2001). The crystal structure of B. subtilis lipase LipA revealed a compact minimal a/b hydrolase fold that lacks a lid domain and a Ca2+-ion as well (van Pouderoyen et al., 2001). Subfamily I.5 lipases originate from different gram-positive bacteria, among them the genera Bacillus, Geobacillus, and Clostridium. The Bacillus lipases included in this family show only little sequence similarity of about 15% to the Bacillus enzymes of family I.4 indicating an evolutionary distant relationship. The pH optima remarkably differ in this subfamily ranging from 9.5 for B. stearothermophilus lipase to 7.5 for the B. thermoleovorans enzyme (Kim et al., 1998; Lee et al., 1999; Rua et al., 1997). The molecular weight of subfamily I.5 lipases is about 46 kDa (Nthangeni et al., 2001) which may be explained by large insertions contained within the canonical a/b hydrolase, e.g., identified in B. stearothermophilus lipase (Tyndall et al., 2002). Here, the insertion forms a zinc-binding site which is unique among all known lipase structures and is believed to provide stability against thermal inactivation (Tyndall et al., 2002). Subfamily I.6 comprises several lipases from Staphylococcus species. They are expressed as preproproteins with a molecular mass of approximately 75 kDa and carry an elongated N-terminal domain of about 200 amino acids that is necessary for efficient translocation and may possibly act as intramolecular chaperone (Goetz et al., 1998; Rosenstein and Goetz, 2000). After secretion, this domain is cleaved off, leaving a mature lipase of 46 kDa molecular mass. These lipases exhibit a wide substrate specificity (Rosenstein and Goetz, 2000; Simons et al., 1998; Tyski et al., 1983) and stability between pH 4 and 9 (Sayari et al., 2001). The activity of some of these lipases was increased by addition of calcium ions and inhibited by EDTA (Nikoleit et al., 1995; Rosenstein and Goetz, 2000; Simons et al., 1999). Interestingly, the activity of Staphylococcus simulans lipase was independent of calcium (Sayari et al., 2001). Staphylococcal lipases tend to form aggregates (Juergens and Huser, 1981; Juergens et al., 1981; Koetting et al., 1983; Vadehra, 1974), and S. simulans lipase forms a tetrameric protein (Sayari et al., 2001). Like many other bacterial lipases, Staphylococcus lipases are virulence factors. S. saprophyticus produces a surface-associated lipase (Sakinc et al., 2005) presumably involved in adhesion to extracellular matrix proteins as collagen or in modification of cellular surface proteins (Shah and Russell, 2004). The lipase from S. hyicus is unique in exhibiting lipase as well as phospholipase activity (Simons et al., 1999; van Oort et al., 1989). Recently, this observation was rationalized from the crystal structure wich revealed that the substrate binding pocket is polar at one side and can thus promote the binding of the phospholipid phosphate group (Tiesinga et al., 2007). Also, this lipase is structurally related to known Geobacillus lipases of family I.5 (Tiesinga et al., 2007). Subfamily I.7 lipases show significant similarity to lipases of family I.2 in the central region of the primary structure comprising amino acid residues 50–150. Two lipases have been identified that belong to this group of enzymes, originating from Streptomyces cinnamoneus (Sommer et al., 1997) and Propionibacterium acnes (Miskin et al., 1997) with molecular masses of 29.2 kDa and 36.4 kDa, respectively. The lipase GehA of P. acnes hydrolyzed a wide range of different substrates including triacylglycerols and p-nitrophenol esters of fatty acids ranging from chain lengths C2–C16 (Falcocchio et al., 2006; Hassing, 1971; Ingham et al., 1981). This observation coincides with its role as a virulence factor which hydrolyzes sebum triacylglycerides (Higaki and Morohashi, 2003) also including a wide range of different lipids and thus mediates the release of inflammatory compounds (Downie et al., 2004).

1109

1110

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Lipolytic Enzymes from Bacteria

Subfamily I.8 represents a new subfamily of lipases which was proposed recently based on the identification and characterization of lipase Lip1 from the psychrophilic bacterium Pseudoalteromonas haloplanktis (de Pascale et al., 2008). Lip1 with 51 kDa molecular mass is associated to the bacterial outer membrane and showed increasing activity against ester substrates with longer acyl chains. Amino acid sequence analysis suggested that this lipase is only distantly related to previously classified lipolytic enzymes; instead, it shows similarity to putative lipases originating from other marine psychrophilic strains like Hahella chejuensis, Colwellia psychrerythraea and Pseudoalterononas tunicata (Holmstrom et al., 1998; Jeong et al., 2005; Methe et al., 2005). Homology modeling revealed that these lipases presumably lack a lid as well as a calcium-binding pocket. Moreover, the sequence LGG(F/L/Y)STG was identified as a novel active site consensus motif replacing the canonical pentapeptide GXSXG surrounding the catalytically active serine (de Pascale et al., 2008).

6

Esterases (Families II–VIII)

Carboxylesterases which hydrolyze water-soluble ester substrates with short chain lengths fatty acids were initially grouped into seven families based on their primary structure and biochemical properties (Arpigny and Jaeger, 1999). Like true lipases, most carboxylesterases belong to the a/b hydrolase fold superfamily of proteins. Exceptions are esterases of family VIII which show a b-lactamase like fold. The BRENDA database (Schomburg et al., 2004) presently contains 696 entries of bacterial carboxylesterases and many of them do not show significant homology to any of the families currently identified (> Table 2).

7

Physiological Functions of Esterases

Esterases can be located either in the cytoplasm (Droege et al., 2005) or in the periplasm (Weadge et al., 2005), be associated to the membrane (von Tigerstrom and Stelmaschuk, 1989; Wilhelm et al., 1999) or secreted (von Tigerstrom and Stelmaschuk, 1989; Xiang et al., 2006). These versatile localizations already suggest manifold physiological functions (see > Fig. 3). The determination of esterase activities has become a valuable tool for monitoring metabolic activity, cell viability, and membrane integrity of bacteria (Amor et al., 2002; Raesaenen et al., 2001; Rhena et al., 2003). As an example, bacterial cells from environmental samples are often in a socalled viable but nonculturable (VBNC) state (Byrd et al., 1991; Roszak and Colwell, 1987; Tanaka et al., 2000). Here, fluorescent substrates are used that enter living cells and are subsequently hydrolyzed by intracellular esterases to yield hydrophilic fluorescent products that accumulate in intact cells allowing their visualization (Tsuji et al., 1995). Moreover, electrophoretic esterase patterns proved useful for phenotyping closely related bacterial strains which are difficult to differentiate, e.g., subspecies of the genera Lactococcus lactis, Pseudomonas aeruginosa and, Escherichia coli (Bert et al., 1995; Gilot and Andre, 1995; Ouzari et al., 2006). Esterases presumably make carbon sources accessible during processes like pathogenic infection, plant cell wall degradation, or detoxification (Khalameyzer et al., 1999). Consistently, esterases are usually not essential for cell survival, at least under standard laboratory conditions (Berger et al., 1998). The probiotic bacterium Lactobacillus reuteri induces the expression of an esterase by a factor of 7.3 upon growth under bile stress, and disruption of the esterase gene results in a residual viability of only 7.5% under these conditions indicating that

Lipolytic Enzymes from Bacteria

21

. Table 2 Families II–VIII of lipolytic enzymes comprising esterases Family II (GDSL)

Origin

Accession No.

Aeromonas hydrophila

P10480

Aeromonas salmonicida

Q44268

Pseudomonas entomophila

Q1IG03

Pseudomonas aeruginosa

O33407

Bacillus thuringiensis

Q3EV80

Pseudomonas sp. HSM0414

Q938A9

Bacillus cereus

Q81AY4

Pseudomonas putida

Q0P6P2, Q6B6R8, A5VXL2

Xanthomonas vesicatoria Q7X4K7 III

IV (HSL)

. . . and 7 others

...

Streptomyces exfoliatus

Q56008

Kineococcus radiotolerans

A6WEQ4

Moraxella sp.

P19833

Clavibacter michiganensis

B0RCM8, B0RFW0

Streptomyces albus

Q59798

Thermobifida fusca

Q47RJ6, Q47RJ7

Alicyclobacillus acidocaldarius

Q7SIG1

Burkholderia sp. 383

Q39FH4, Q39NZ4, Q38ZY5, Q390B6, Q397N4, Q39AV1, Q39B99, Q39NZ2

Archaeoglobus fulgidus

O28558

Bradyrhizobium sp. ORS278

A4YTP9

Ralstonia eutropha

Q44019

Pseudomonas sp. B11–1

O52270

Bradyrhizobium sp. BTAi1 A5EJD5, A5EAZ6 Ralstonia solanacearum

A3RSM7

Bacillus subtilis

O68884

Bacillus thuringiensis

Q3EL41, Q3EX93

. . . and 19 others

...

1111

1112

21

Lipolytic Enzymes from Bacteria

. Table 2 (Continued) Family V

Origin

Accession No.

Psychrobacter immobilis

Q02104

Moraxella sp.

P24640

Salmonella typhimurium

Q8ZNL5

Rhodoferax ferrireducens

Q223C0

Enterobacter sp. 638

A4WCJ0

Serratia proteamaculans

A8GC20

Polaromonas naphthalenivorans

A1VLL6

Rhodobacter sphaeroides A4WX93, A3PMP7 VI

. . . and 85 others

...

Spirulina platensis

Q53415

Anabaena variabilis

Q3M6G8

Bordetella avium

Q2KUZ2

Polaromonas sp. JS666

Q127I1

Shewanella amazonensis

A1S771

Xanthomonas campestris B0RNI6, Q3BXV6

VII

VIII

Rhodoferax ferrireducens

Q21XU9

. . . and 49 others

...

Arthrobacter oxydans

Q01470

Bacillus stearothermophilus

Q8GCC7

Bacillus sp.

Q9X6Z3

Bacillus sp. BP-7

Q9L378

Bacillus amyloliquefaciens

A7Z924

Streptomyces coelicolor

Q9Z545

Burkholderia sp. 383

Q398P3

Bacillus pumilus

Q66M67

. . . and 15 others

...

Arthrobacter globiformis

Q44050

Arthrobacter aurescens

A1RB78

Saccharopolyspora erythraea

A4F8E6

Pseudomonas fluorescens Q8VU79, Q4KH73 Pseudomonas gingeri

B0M0H4

Pseudomonas syringae

Q48LQ9

Streptomyces anulatus

O87861

Lipolytic Enzymes from Bacteria

21

this esterase may be involved in reorganization of the cellular membrane (Wall et al., 2007; Whitehead et al., 2008). Additionally, it was shown that increasing H2O2 concentrations induced alterations of the esterase expression levels in four different bacterial strains implying that these enzymes aid the bacteria to cope with oxidative stress (Baatout et al., 2006). Bacterial esterases are also important as regulators of interspecies communication and interaction. Several esterases degrade quorum-sensing (QS) signal molecules, e.g. an enzyme from Ideonella species which can degrade 3-hydroxypalmitic acid methyl-ester (3-OH PAME), the QS signal of the plant pathogen Ralstonia solanacearum (Shinohara et al., 2007). Pseudomonas fluorescens esterase degrades lactones (Khalameyzer et al., 1999) which serve as QS signals in many bacterial species including Pseudomonas. In Streptomyces species, esterases hydrolyze storage triacylglycerols and the resulting fatty acids serve as carbon sources for antibiotic synthesis (Olukoshi and Packter, 1994). Another interesting observation was made by identifying two esterases in a metagenomic library. In both cases, these enzyme genes flanked orf ’s which encode enzymes related to antibiotic resistance, like a b-lactamase and a putative aminoglycoside transferase (Elend et al., 2006). Esterases, much like lipases, are related to the virulence of pathogenic bacteria. Plant cell wall degrading bacteria, among them Clostridium and Lactobacillus species (Donaghy et al., 2000; Prates et al., 2001; Wang et al., 2004) can hydrolyze ester bonds between hydroxycinnamic acid and sugars (Garcia-Conesa et al., 1999). Human pathogenic esterases include EstV from Helicobacter pylori which is directly linked to the development of peptic ulcers and stomach cancer (Kusters et al., 2006) and represents the only lipolytic enzyme identified in this organism (Ruiz et al., 2007). This pathogen can degrade the lipids of the gastric mucus (Slomiany et al., 1989), most likely by the activity of EstV. The nosocomial pathogen Corynebacterium jeikeium, which causes septicaemia and endocarditis, lacks genes encoding fatty acid synthase (Hansmeier et al., 2007). Instead of biosynthesizing fatty acids, this bacterium produces a cholesterol esterase which can hydrolyze fatty acid esters from cholesterol present in the blood and in extracellular matrix proteins (Hansmeier et al., 2007; Tauch et al., 2005). The pathogenicity of Mycobacterium tuberculosis strongly depends on lipolytic enzymes as well. Attenuated M. tuberculosis strains constructed by transposon mutagenesis revealed that most of the mutated loci were involved in metabolism or membrane transport of lipids (Camacho et al., 1999), among them the gene encoding LipF which belongs to esterase family IV (Zhang et al., 2005). In Pseudomonas aeruginosa, the esterase EstA was identified as an autotransporter which may also be involved in virulence. An estA deletion mutant was strongly impaired in cell motility as well as rhamnolipid production and the formation of biofilms (Wilhelm et al., 2007).

8

Family II

This family comprises the so-called GDSL/SGNH enzymes which lack the conserved lipase active site motif G-X-S-X-G, and instead contain a G-D-S-(L) tetrapeptide located in the N-terminal part of the proteins (Arpigny and Jaeger, 1999). The GDSL family of hydrolases was identified in 1995 (Upton and Buckley, 1995) and contains five blocks of conserved amino acid residues with the above mentioned tetrapeptide located in block 1. Another family of hydrolases was proposed on the basis of four protein crystal structures including a rhamnogalacturonan acetylesterase of Aspergillus aculeatus, the esterases of Streptomyces scabies and influenza C virus and an acetylhydrolase originating from Bos taurus (Molgaard et al., 2000;

1113

1114

21

Lipolytic Enzymes from Bacteria

Molgaard, 2002). The enzymes belonging to this family show significant homology in four blocks of their primary structure, and one strictly conserved amino acid was identified in each block which is essential for catalysis, namely serine, glycine, asparagine, and histidine. Thus, these enzymes were referred to as SGNH hydrolases. The first block of conserved amino acids carries the catalytically active serine located within the GDS(L) motif (Molgaard, 2002). The Gly conserved in block II and the Asn in block III serve as proton donors to the oxyanion hole. The basic His located in block V increases the nucleophilicity of the Ser by deprotonating its hydroxyl group. A comparison of the classical a/b hydrolase fold with the fold of SGNH hydrolases revealed a different location of the residues involved in active side formation resulting in a different relative orientation of the catalytic triad with regard to the central parallel b sheet. While in a/b hydrolases, the catalytic triad aligns parallel to the sheet, in SGNH hydrolases, an almost perpendicular alignment could be identified (Molgaard et al., 2000). Some GDSL hydrolases, e.g. from P. aeruginosa, Salmonella typhimurium and Photobacterium luminescens share an additional domain located at the C-terminus of the respective proteins and comprising approximately one third of the entire protein. This domain consists of 12 b-sheets which form a b-barrel inserted into the bacterial outer membrane, similar to the structures of porines (Henderson et al., 1998; Oomen et al., 2004; Tamm et al., 2001). The N-terminal part of these proteins which harbors an enzymatic activity is exported through this b-barrel and is often cleaved off after translocation to release the N-terminal enzyme domain (Dautin and Bernstein, 2007; O’Toole et al., 1994; Pohlner et al., 1987; St Geme and Cutter, 2000; Steinhauer et al., 1999; Suhr et al., 1996). These proteins were termed autotransporters (Loveless and Saier, 1997) and have since been identified in several pathogenic bacteria where they contribute to virulence (Ma et al., 2003; Rosenau and Jaeger, 2000; Wilhelm et al., 1999).

9

Family III

Extracellular enzymes from Streptomyces and Moraxella species (Cruz et al., 1994; Feller et al., 1990; Perez et al., 1993) belong to this family. Additionally, hydrolases of Acidovorax and Thermobifida species were identified displaying high levels of similarity to these lipases and being able to degrade polyesters (Kleeberg et al., 2005; Uchida et al., 2002). Lipases of family III have a molecular mass of 32–35 kDa and show significant similarity to the intracellular and plasma monomeric isoforms of the human platelet activating-factor acetylhydrolase (PAF-AH) (Arpigny and Jaeger, 1999). The crystal structure of a lipase from S. exfoliatus showed a canonical a/b hydrolase fold containing the conserved catalytic triad, but lacked a lid as found in the PAF-AHs (Wei et al., 1998). In S. coelicolor, a growth-phase dependent regulation of lipase expression was identified with the highest level of enzyme synthesis occurring at the stationary phase (Servin-Gonzalez et al., 1997). A specific activator protein designated LipR was identified which is encoded in an operon together with the lipase and binds upstream of the lipase promoter inducing transcription (Evangelista-Martinez et al., 2006; Servin-Gonzalez et al., 1997).

10

Family IV

This family constitutes several esterases from distantly related prokaryotes including psychrophilic, mesophilic, and thermophilic bacteria. The enzymes show a remarkable similarity to

Lipolytic Enzymes from Bacteria

21

the family of mammalian hormone-sensitive lipases (HSL) (Hemilae et al., 1994). Here, three sequence blocks with conserved motifs were identified, with blocks II and III containing the catalytic triad residues (Arpigny and Jaeger, 1999). Block I contains a conserved H-G-G-G consensus sequence that is involved in hydrogen bonding interactions which stabilize the oxyanion hole and promote catalysis as deduced from the crystal structure of Brefeldin A esterase from Bacillus subtilis (de Simone et al., 2000; Manco et al., 1999; Wei et al., 1999). Despite their homology to HSL, the substrate spectra of the enzymes of the bacterial HSLfamily differ significantly. While human HSL has a broad substrate specificity and hydrolyzes short chain esters as well as water insoluble substrates like trioctanoin, vinyl laurate and olive oil, the bacterial HSL-family esterases show activity only towards short fatty acid chain length substrates like tributyrin and vinyl propionate (Chahinian et al., 2005). Moreover, esterase enzyme kinetics were observed while HSL reactions show typical lipase kinetics (Ben Ali et al., 2004; Chahinian et al., 2005). Several family IV esterase structure were solved (de Simone et al., 2000; de Simone et al., 2001; Wei et al., 1999; Zhu et al., 2003) which showed a unique structural feature: a ‘‘cap’’ which clearly differs from the lid found in true lipases (Wang et al., 2005) is formed by two separate helical regions and covers the active site. More recently, an oxadiazolone inhibitor was identified which covalently binds to the catalytically active serine. Interestingly, inhibition was specific in that the activity of other carboxylesterases was not affected providing a promising option for quick discrimination among esterases (Ben Ali et al., 2006).

11

Family V

Esterases of this family originate from different bacterial genera representing mesophilic, cold-, or heat-adapted organisms as Pseudomonas, Haemophilus, and Moraxella. These enzymes share significant homology with other bacterial enzymes like epoxide hydrolases, dehalogenases, and haloperoxidases that also possess the typical a/b hydrolase fold (Arpigny and Jaeger, 1999). Conserved amino acids are located in three blocks with the catalytic triad residues Ser located in block II, Asp, and His in block III. The esterases Est2 of Acetobacter pasteurinanus and EstVof Helicobacter pylori are the only enzymes of this family that have been cloned and characterized yet (Kashima et al., 1998, 1999; Ruiz et al., 2007). Both enzymes revealed typical characteristics of carboxylesterases with EstV of H. pylori showing preference for short fatty acid chain length p-nitrophenyl esters (C2–C6) and triglycerides (C4) and typical Michaelis-Menten kinetics instead of interfacial activation (Ruiz et al., 2007). In case of A. pasteurianus Est2, triglyceride substrates with even shorter fatty acid chain lengths like triacetin and tripropionin were hydrolyzed preferably (Kashima et al., 1999).

12

Family VI

Enzymes classified in family VI are small proteins ranging from 23 to 26 kDa, with the exception of an esterase isolated from a metagenomic library which had a molecular mass of 31.6 kDa (Arpigny and Jaeger, 1999; Kim et al., 2006). These enzymes exhibit a sequence similarity of about 40% to eukaryotic lysophospholipases and three conserved blocks, as already found in families IV and V, were identified (Arpigny and Jaeger, 1999). A carboxylesterase from Pseudomonas fluorescens was the first member of this family to be cloned, expressed, and characterized. The native enzyme forms a dimer and shows activity

1115

1116

21

Lipolytic Enzymes from Bacteria

against short fatty acid chain lengths substrates like p-nitrophenylacetate and p-nitrophenylbutyrate (Hong et al., 1991). A carboxylesterase of P. aeruginosa showed a similar substrate spectrum: it hydrolyzed preferably short chain substrates like p-nitrophenyl acetate and moreover 4-methyl-umbelliferyl acetate but did not accept triacylglyceride substrates (Pesaresi et al., 2005). Recently, a novel esterase originating from Pseudomonas sp. CR-611 was cloned and characterized which showed maximum activity at 55 C and pH 6.5 (Prim et al., 2006). A dimeric quarternary structure was observed in crystal structures of Pseudomonas fluorescens esterase and Pseudomonas aeruginosa PAO1 carboxylesterase (accession number PA3859) (Kim et al., 1997; Pesaresi and Lamba, 2005). Subunit dimerization involves three active-site loops of each monomer (Kim et al., 1997). The physiological functions of bacterial family VI esterases still remain elusive, but it is known for human acyl-protein thioesterase I which shows significant sequence and structure similarity to members of this esterase family. Its in vivo function is the S-palmitoylation of cysteine residues of G protein alpha subunits. (Devedjiev et al., 2000; Duncan and Gilman, 1998; Pesaresi et al., 2005).

13

Family VII

Esterases belonging to family VII have a molecular mass of approximately 55 kDa and show high similarity to acetylcholine esterases and intestine/liver carboxylesterases (Arpigny and Jaeger, 1999). Enzymes originating from several Bacillus, Peanibacillus, and Geobacillus strains are found in this group. Four blocks of significantly conserved amino acids were identified allowing to design degenerated primers and amplify family VII esterase homologous genes from other Bacillus genomes as well as from soil and mine water samples (Nthangeni et al., 2005). Typical carboxylesterase substrates hydrolyzed by these enzymes include short fatty acid chain length p-nitrophenyl esters and triglycerides, but enzymes of this family also hydrolyzed biotechnologically relevant substrates. The esterase PnbA from B. subtilis hydrolyzed p-nitrobenzyl esters and may thus be used to remove protecting groups during the synthesis of b-lactam antibiotics (Zock et al., 1994). An esterase of B. subtilis strain RRL BB1 enantioselectively hydrolyzed racemic 2-hydroxy-4-phenylbutanoate to yield the (S)-enantiomer (ee  80%) which serves as an intermediate in the synthesis of inhibitors of angiotensinconverting enzymes (Maqbool et al., 2002, 2006). Another esterase derived from Arthrobacter oxydans showed activity against phenylcarbamate herbicides like phenmedipham. Since this xenobiotic compound has no equivalent in nature, the physiological substrate for this enzyme is still unknown (Pohlenz et al., 1992).

14

Family VIII

In contrast to all other lipases and esterases characterized so far, the structures of family VIII enzymes differ significantly from the typical a/b-hydrolase fold and instead show remarkable similarities to b-lactamases as well as DD-peptidases which catalyze cross-linking of D-amino acids in bacterial peptidoglycan. A prototype enzyme is the esterase EstB from Burkholderia gladioli (formerly named Pseudomonas marginata) (McKay et al., 1992; Wagner et al., 2002)

Lipolytic Enzymes from Bacteria

21

which consists of a b-sheet core surrounded by a-helices, however, the core is formed primarily by antiparallel b-strands. Moreover, the catalytically active serine is not part of a catalytic triad, and it is located at the beginning of an a-helix in the vicinity of the central sheet (Wagner et al., 2002). Site-directed mutagenesis studies and the crystal structure of B. gladioli esterase solved in complex with an inhibitor clearly demonstrated that the nucleophilic serine was located in a S-X-X-K motif rather than within one of the lipase consensus motifs G-X-SX-G or GDSL (Petersen et al., 2001; Wagner et al., 2002). Additional to the serine and the lysine residues of the N-terminal located consensus motif, a tyrosine was identified as being essential for enzymatic activity (Sakai et al., 1999). Interestingly, an additional G-X-S-X-G motif was identified as well which was located in the C-terminal part of these esterases (Kim et al., 1994; Petersen et al., 2001). Esterases belonging to family VIII have a molecular mass of approximately 42 kDa, and for some of them, multimerisation appeared necessary to form enzymatically active enzymes (Elend et al., 2006; Nishizawa et al., 1995; Schuette and Fetzner, 2007), while others were monomers (Kim et al., 1994; McKay et al., 1992; Rashamuse et al., 2007; Wagner et al., 2002). The high solvent tolerance of some of these esterases may represent a biotechnologically interesting feature (Elend et al., 2006; Ogino et al., 2004; Schuette and Fetzner, 2007). Esterases of family VIII and b-lactamases are structurally related, but reactivity towards both classes of substrates was reported only occasionally (Govardhan and Pratt, 1987; Jones and Page, 1991; Pratt and Govardhan, 1984). Esterase EstB from B. gladioli exhibited deacetylation activity on cephalosporin derivates only, but b-lactamase activity was not detectable for this and for other esterases (Petersen et al., 2001; Rashamuse et al., 2007; Sakai et al., 1999).

15

Research Needs and Conclusions

At present, about 1,500 lipolytic enzymes have been identified with almost 900 of them originating from bacteria (> Fig. 2). It should be noted that most of these enzymes just represent orfs annotated within numerous genome and metagenome sequencing projects. Nevertheless, true lipases and carboxylesterases comprise a huge diversity in terms of molecular mass, structure, and biochemical properties. They are widespread throughout different bacterial strains including gram-positive, gram-negative, and cyanobacteria as well as archaea. Unfortunately, the physiological role of many of these enzymes still remains elusive, however, they obviously carry out a variety of cellular and extracellular functions other than just providing carbon sources by hydrolysis of lipidic substrates (> Fig. 3). This notion is substantiated by the complexity of gene regulation mechanisms and the diversity of different cellular localizations observed for these lipolytic enzymes. It is not a surprise that some lipolytic enzymes cannot be classified into currently existing enzyme families clearly suggesting that more families may exist. Indeed, a new family of lipolytic enzymes has recently been suggested (de Pascale et al., 2008). Also, a lipase originating from Thermotoga maritima was described which displays high amino acid sequence similarity to subfamily I.2 enzymes, however, the region around the conserved pentapeptide containing the catalytic serine exhibits closer similarity to family I.4 lipases (Kakugawa et al., 2007). Undoubtedly, research on lipolytic enzymes remains a fascinating field with many new and exciting enzymes to be discovered.

1117

1118

21

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Part 5

Genetics (the Paradigms) Section Editor: Victor de Lorenzo

22 Transcriptional Control of the TOL Plasmid Pathways P. Domı´nguez-Cuevas . S. Marque´s* Department of Environmental Protection, Estacio´n Experimental del Zaidı´n, Granada, Spain *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1130 2 The Upper Pathway: Onset of the Regulatory Cascade . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1131 3 The Meta-Cleavage Pathway: New Insights into the Activation Mechanism . . . . . . . 1133 4 Integration in the Cell Regulatory Networks: Toward Optimization of Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1134 5 Research Needs and Future Perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1136

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_78, # Springer-Verlag Berlin Heidelberg, 2010

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Transcriptional Control of the TOL Plasmid Pathways

Abstract: The TOL plasmid encoded pathway for the degradation of toluene and derivatives is an archetype in bacterial transcription regulation. Six promoters of different type and several chromosome and plasmid-encoded proteins are involved in maintaining optimal expression levels and synchronization with the global cell metabolism. The TOL- encoded regulators are the enhancer binding protein XylR, which controls the upper pathway Pu and xylS gene PS1 s54-dependent promoters, and the AraC family regulator XylS, which controls the s32 s38-dependent meta-cleavage pathway promoter Pm. Both regulators respond to the presence of a specific effector and activate transcription through different mechanisms. Much effort has been devoted to the elucidation of these processes. In this review, recent results are described and discussed in the light of the latest findings and models for homologous family proteins.

1

Introduction

Pseudomonas putida mt-2 TOL plasmid pWW0 was one of the first plasmids described to contain an entire aromatic degradation pathway (Williams and Murray, 1974). It codes for two catabolic routes responsible for toluene and benzoate degradation, the upper and meta-cleavage pathways, respectively (Assinder and Williams, 1990), flanked by two insertion sequences (IS1246). The genes are organized in two operons located 10-kb apart (upper and meta-cleavage), clustered next to the divergent regulatory genes xylR and xylS, which coordinate the expression of the two pathways (Greated et al., 2002; Harayama and Timmis, 1989). The regulatory network can be described as follows: when P. putida mt-2 cultures are exposed to toluene, the XylR regulator becomes active to promote transcription from two s54-dependent promoters: the upper pathway promoter Pu and the xylS gene promoter PS1 (Delgado and Ramos, 1994; Dixon, 1986; Gallegos et al., 1996a; Inouye et al., 1987; Marque´s et al., 1998; Ramos et al., 1987). Activation of PS1 leads to increased synthesis of XylS, which is able to promote expression from the meta-cleavage pathway promoter Pm even in the absence of its effector. This mechanism is known as the cascade regulatory loop (Marque´s and Ramos, 1993; Ramos et al., 1997). In the absence of toluene, transcription from Pm can be induced by the addition of benzoate and some derivatives to activate XylS, a regulatory pathway known as meta-loop (Inouye et al., 1987; Ramos et al., 1986). On the other hand, toluene-dependent XylR induction of the upper pathway also leads to the metabolism of toluene to benzoate, the first substrate of the meta-cleavage pathway. In turn, this aromatic activates the XylS regulator to promote increased transcription from the meta-cleavage pathway promoter Pm and the resulting synthesis of pathway enzymes. The extensive analysis of these processes shows that a considerable number of chromosome and plasmid encoded proteins cooperates to maintain an optimal pathway expression level in every circumstance (> Table 1). The intricacy and refinement of the regulatory network allow for an appropriate synchronization with the global cell metabolism, and makes of the TOL system an archetype in bacterial transcription regulation. The general patterns of this network have already been revised (Daniels et al., 2008; Marque´s and Ramos, 1993; Ramos et al., 1997; Ruı´z et al., 2004). This chapter focuses on the most recent findings, especially those explaining the mechanistic and fine-tuned functioning of the pathway and its integration into the cell regulatory network.

Transcriptional Control of the TOL Plasmid Pathways

22

. Table 1 Protein factors involved in TOL pathway regulation Factor

Gene(s)

Category

Target/rolea

Sigma 70

rpoD

General, house keeping

xylR (+), xylS (+)

Sigma 38

rpoS

General, stress/stationary

meta-cleavage pathway (+)

Sigma 32

rpoH

General, Stress

meta-cleavage pathway (+)

Sigma 54

rpoN

General, nitrogen, other functions

Upper pathway (+), xylS (+)

IHF

himA, hip

General

Upper pathway (+), xylS ( )

IIA

ptsN

General

Upper pathway, xylS

Crc

Crc

General

Upper pathway ( )

FtsH

ftsH

General

Upper pathway (+)

XylR

xylR

Pathway specific

Upper pathway (+), xylS (+)

XylS

xylS

Pathway specific

meta-cleavage pathway (+)

Ntr

a

(+) positive effect; ( ) negative effect

2

The Upper Pathway: Onset of the Regulatory Cascade

XylR is the primary regulator of the TOL pathway. It belongs to the large AAA+ family of ATPases (Neuwald et al., 1999; Studholme and Dixon, 2003) which includes, among others, the s54-dependent promoter activator family known as enhancer binding proteins (EBPs). These proteins consist of three main domains: an N-terminal input domain (A) sensing the regulatory signal (e.g., the presence of an effector in the case of XylR (Delgado and Ramos, 1994)), a central AAA+ domain (C) with ATPase activity, and a C-terminal DNA-binding domain(D) (Studholme and Dixon, 2003). The role of the three domains has been well established in XylR. Genetic analysis located the mutations altering the effector profile in the N-terminal domain of the protein (Delgado and Ramos, 1994; Garmendia et al., 2001, 2008; Salto et al., 1998). This domain given in trans exerted specific intramolecular repression inhibiting ATP binding capacity of the C domain, but repression was released in the presence of an effector (Ferna´ndez et al., 1995; Pe´rez-Martı´n and De Lorenzo, 1995). Bioinformatic analysis based on the alignment of 11 XylR family proteins predicts the eight a-helix and seven b-strands shape of this domain structure, leaving the neutral effector binding pocket in a shallow groove in the domain surface (Devos et al., 2002). Although in XylR the physical interaction with the effector has not yet been analyzed, effector binding analysis in the phenolresponsive XylR homolog DmpR revealed that the interaction of labeled phenol with the Nterminal domain released C-domain ATPase activity repression (Shingler and Pavel, 1995). The A domain is connected to the central domain by a short coiled-coil structure linker (B) which has been suggested to influence effector binding and to be involved in protein oligomerization (Garmendia and de Lorenzo, 2000). Recent analyses of EBP structures show that this domain regulates multimerization, and couples the A-domain input signal with C-domain ATPase activity (Bose et al., 2008). Although this has not been directly demonstrated in XylR, the strong homology among these proteins points toward a similar mechanism operating in the TOL regulator.

1131

1132

22

Transcriptional Control of the TOL Plasmid Pathways

The AAA+ central domain (C) is the most conserved among EBPs. It carries the ATP binding motif, ATPase activity and s54 interaction determinants, features that are essential to s54 promoter activation (Chen et al., 2008; Schumacher et al., 2006). According to the first XylR activation model based on the analysis of a truncated protein devoid of its N-terminal domain, the activation mechanism followed a cyclic sequence of events where ATP binding to the central domain triggered XylR multimerization at its binding site, followed by ATP hydrolysis, promoter activation and return to the non-multimerized structure (Pe´rez-Martı´n and de Lorenzo, 1996). The recent availability of the crystal structure of several regulators of the family has helped to increase our knowledge of the multistep process leading to promoter activation by these proteins (Lee et al., 2003; Rappas et al., 2005; Sallai and Tucker, 2005). Although the mechanistic model has only been proposed for those members belonging to a two component signal, XylR is likely to share many of its features. Proteins of this family generally bind DNA as dimers. After the A domain repression has been released (in XylR, by effector binding), a conformational rearrangement induces protein oligomerization to a DNA-bound hexameric structure, which is followed by ATP binding to the regulator in the interface of two subunits. Interestingly, both adjacent subunits contribute to ATP hydrolysis, thus explaining the previously observed ATPase dependence on protein oligomerization (Bordes et al., 2003). This feature has been further reviewed by (Schumacher et al., 2006). Two C-domain EBP specific loops pointing toward the inner pore of the hexameric ring undergo a conformational reorganization during ATP hydrolysis, reorienting the AAA-family conserved GAFTGA motif present in one of the loops to allow contacts with s54 (Bose et al., 2008; Wigneshweraraj et al., 2008). Contacts between the regulator and the transcriptional machinery require the two proteins are brought into proximity by DNA bending, generally facilitated by IHF. Finally, s54 undergoes a structural remodeling, allowing Es54 closed complex to isomerize to open complex. In the protein sequence, the central domain is followed by the DNA-binding domain (D) encompassing a typical helix-turn-helix (HTH) structure which confers binding specificity on the target promoter (Inouye et al., 1988). In Pu XylR dimers are always bound to the two upstream activation sequences (UAS) (Abril et al., 1991). As shown above for other EBPs, binding of ATP induces multimerization to an hexameric conformation, which is then able to hydrolyze ATP (Pe´rez-Martı´n and de Lorenzo, 1996). The relevant regulatory sequences in Pu span 108 bp and the XylR binding site is composed of two UASs located between positions 120 and 175 (Holtel et al., 1990). Interestingly these UASs, the furthermost sequence of the TOL xyl region located adjacent to IS1246, prevent read-through from upstream promoters, thus isolating pathway expression from external influence (Vela´zquez et al., 2006). UV-laser footprint and atomic force microscopy confirmed binding of IHF between positions 52 and 79 (Valls et al., 2002), inducing the strong DNA bending needed for interaction between the UAS-bound regulator and the RNAP machinery bound at 12/ 24 (de Lorenzo et al., 1991). Early models explaining EBP activation mechanism suggested direct interaction between the regulator and the RNAP, in most cases brought to proximity by IHF assisted DNA looping. Although this is essentially true, accumulated evidence in recent years and the availability of crystal structures show that surprisingly, contacts between regulator and RNAP are established with an unexpected orientation. In fact, the regulator approaches the RNAP closed complex from the unbound face of RNAP binding site to contact s54 and catalyzes open complex formation, so that DNA appears sandwiched between RNAP and regulator (Huo et al., 2006). The precise mechanism underlying energy coupling from regulator ATP hydrolysis to promoter melting and escape still remains unknown.

Transcriptional Control of the TOL Plasmid Pathways

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The structure of the PS1 promoter slightly differs from the canonical s54-promoter architecture, probably because it accommodates the two overlapping divergent promoters PR1 and PR2 responsible for xylR expression, and PS2 xylS constitutive promoter which maintains XylS basal concentrations in the cell (Gallegos et al., 1996a). In fact, XylR UASs at position 133 to 207 of PS1 overlap the divergent 10/ 35 s70 RNAP binding sites of the two xylR promoters. Two consensus IHF binding sequences are found, overlapping the 12/ 24 s54 RNAP binding site and the UASs. As a consequence of this complex organization, regulator expression never achieves maximum levels (Marque´s et al., 1998): when XylR binds its UASs to activate PS1, it represses its own synthesis (Bertoni et al., 1998; Holtel et al., 1992); and surprisingly, the presence of IHF strongly represses PS1 activity.

3

The Meta-Cleavage Pathway: New Insights into the Activation Mechanism

Expression of the meta-cleavage pathway is under the control of XylS-regulated Pm promoter. Expression along the growth curve is mediated by two stress sigma factors: s32 in the early exponential phase and s38 in the late exponential and stationary phases (Marque´s et al., 1999). In fact, Pm 10/ 35 RNAP binding sequence diverges considerably from the consensus defined for s32, s38 and s70 factors (Domı´nguez-Cuevas et al., 2005). On one hand XylS binding sites overlap the 35 region so this sequence is a compromise between the two binding consensus (Gonza´lez-Pe´rez et al., 2002); on the other hand the 10 region of Pm must include the essential determinants for recognition by the two polymerases involved, Es32 and Es38. Unlike s70, the amount of these two alternative sigma factors depends on the cell physiological state, and requires stress conditions to reach effective amounts to compete with s70 for core RNAP (Gross et al., 1998; Hengge-Aronis, 2002). Global expression analyses of P. putida mt-2 revealed aromatic effectors such as toluene or 3-methylbenzoate (3MB) are good elicitors of the stress response (Domı´nguez-Cuevas et al., 2006). A thorough mutational analysis of Pm 10 region showed that s32 and s38 do not share many recognition elements in the promoter. A GC at positions 9 and 8 and the CCCC sequence upstream from the 10 element were crucial for Es32 recognition. Position 10 was the only essential one for s38 recognition, and GC at 8/ 9 prevented Es38 competition for Pm binding (Domı´nguez-Cuevas et al., 2005). This general picture suggests the Pm promoter sequence has evolved to adapt the meta-cleavage pathway expression to the heat shock response triggered by the presence of toluene or 3MB (Domı´nguez-Cuevas et al., 2006). The increase in s32 level during the heat-shock response is strong but transient, so the Pm promoter sequence is optimized to accommodate the temporary Es32 RNAP. After this initial heat shock response, a second alternative RNAP associated with the stress/stationary sigma factor s38 maintains Pm transcription at high levels. Thus the roles of effector in Pm promoter activation mechanism are to activate XylS protein and to increase stress sigma factor levels to efficiently compete for core binding and promoter recognition. XylS binding site at Pm is composed of two 15-bp direct repeats (positions 70 to 56 and 49 to 35) each divided in two sequence boxes A and B. Pm exhibits an intrinsic curvature centered in the A-track located between proximal boxes A and B, with an apparent bent angle of 35 , also observed in vivo (Gallegos et al., 1996b; Gonza´lez-Pe´rez et al., 2002). XylS belongs to the AraC family of transcriptional regulators (Gallegos et al., 1997; Tobes and Ramos, 2002) and is composed of two separate functional domains: XylS mutants

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with altered effector specificity cluster in the N-terminal domain of the protein, indicating this domain carries the effector recognition determinants (Micha´n et al., 1992; Ramos et al., 1990; Ruı´z and Ramos, 2002). In addition, mutations in residues Leu193, Leu194 and Ile205 on the C-terminal edge of this domain are impaired in XylS dimerization (Ruı´z et al., 2003). On the other hand, genetic analysis located the DNA binding domain at the C-terminal end of the protein, connected to the N-terminal domain by a short linker. X-ray crystallographic structures of other family proteins show that the DNA-binding domain is composed of seven a-helices folding in two HTH domains, which bind two adjacent segments of the major groove (Kwon et al., 2000; Rhee et al., 1998). XylS domains are functionally independent, so that XylS DNA-binding domain (XylSC) is able to activate transcription in spite of the absence of XylS N-terminal domain determinants, a process independent of the presence of effector (Domı´nguez-Cuevas et al., 2008b). Analysis of this truncated protein showed it bound DNA forming two complexes (CI and CII), corresponding to one or two XylS-C monomers bound to DNA, respectively (unpublished). Affinity calculation, DNA bending angle estimation and footprinting assays of XylS C-terminal domain suggest the two monomers bind DNA cooperatively, so that when the first XylS-C monomer binds Pm at the proximal site (closest to the RNAP binding site), Pm curvature raises from 35 to 50 . Simultaneously, the bent center shifts to the DNA region between XylS binding sites and finally the binding of the second XylS-C monomer increases the DNA bending angle to 98 . This probably contributes to establish the XylS-RNAP contacts required for transcription activation. Our results indicate that sugarphosphate backbone contacts greatly contribute to XylS/Pm binding strength (Domı´nguezCuevas et al., 2008b), so that Pm curvature around XylS monomers probably enhances nucleoprotein stability. In addition, XylS establishes base specific contacts that are on the basis of unambiguous recognition of Pm direct repeats (Domı´nguez-Cuevas et al., 2008b). XylS dimer formation and DNA binding capacity were enhanced in vivo by the presence of 3MB, but became an effector independent process at high protein concentration in vitro (Domı´nguez-Cuevas et al., 2008a; Ruı´z et al., 2003). As for the role of 3MB in XylS activation, data obtained with the two purified protein domains suggest intramolecular repression of XylS-N upon XylS-C DNA binding, which was released in the presence of 3MB (Domı´nguezCuevas et al., 2008a). The current model for XylS activation involves the following sequence of events: in the absence of 3MB, direct interaction between N- and C-terminal domains in XylS maintains the protein in an inactive state. The addition of 3MB releases N-terminal domain repression upon XylS-C, allowing XylS to bind DNA and to activate transcription. Thus 3MB binding to XylS both triggers the conformational change favoring the dimerization and allows de-repression of the DNA binding domain. However, a XylS dimerization mutant able to bind DNA in the presence of 3MB remained inactive in transcription, indicating dimerization is an essential process in transcription activation. The mode of activation is slightly different in the absence of effector: at high XylS concentrations XylS C-terminal domain de-repression is favored, unmasking dimerization surface and DNA binding determinants, which leads to Pm recognition and further activation.

4

Integration in the Cell Regulatory Networks: Toward Optimization of Expression

As listed in > Table 1, TOL-mediated degradation of aromatic compounds involves a number of cell global regulators. This denotes a long coexistence of plasmid and host, leading to the

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adaptation of plasmid gene expression to the cellular metabolism, taking advantage of the host machinery and using its network of general regulators. The main targets of this network are the s54-dependent Pu and PS1 promoters, and Pm. Pu and PS1 respond to the availability of alternative carbon source, while Pm is optimized to maintain significant expression levels under induced conditions. Factors affecting Pm expression have already been addressed above. Influence of global physiological conditions on Pu expression is observed in rich medium as a delay to induce the pathway after effector addition, which has also been called ‘‘exponential silencing’’ (reviewed in (Cases and de Lorenzo, 2005)), and in defined medium as reduced activation levels when, as well as the effector, an additional carbon source such as glucose, gluconate or 2-ketoglutarate is present (reviewed in (Ruı´z et al., 2004)). These effects are mimicked in continuous culture, in which growth under carbon excess leads to total repression of the pathway regardless of the carbon source used and regardless of the limiting substrate selected, while growth in carbon-limited conditions allows substantial expression (Duetz et al., 1996). The mechanisms underlying this modulation have been analyzed for years and a number of factors involved in the process have been described (> Table 1). However, no clear picture of the regulatory network has emerged yet. The first unambiguous outcome from these analyses was that the classical CRP-dependent regulation present in Enterobacteriaceae is not involved in this P. putida regulation (Suh et al., 2002). A summary of the latest physiological findings in TOL global regulation can be found in (Daniels et al., 2008) (this volume). Three additional targets for this global regulation of the TOL system have been considered. The first one was based on the regulatory network prevailing in Pseudomonas CF600 for the homologous system DmpR/Pdmp, which involves the alarmone (p)ppGpp as main player in global regulation (Sze and Shingler, 1999). However, it seems this mechanism has little influence on XylR-mediated Pu expression (Carmona et al., 2000). The sigma factor s54 was also explored as possible target for global regulation. In most Gram negative bacteria where the rpoN gene coding for this factor is found, it appears clustered with three other genes: ptsN, which codes for the phosphoenolpiruvate:sugar phosphotransferase system (PTS) component IIANtr (Deutscher et al., 2006), is not connected to any specific sugar intake and is suggested to control carbon/nitrogen balance in the cell (Cases et al., 2007). The two additional C/N PTS enzyme genes are ptsP (EINtr) and ptsO (NPr). The phospho-relay cascade of this PTS system flows from phosphoenol pyruvate (PEP) through EINtr, NPr to IIANtr. Surprisingly, in P. putida this pathway is interconnected with the fructose transport system (Pfluger and de Lorenzo, 2008), leading to the suggestion that carbon flux through the Entner–Doudoroff pathway would be translated to the C/N PTS system through FruB. However, the final target of IIANtr regulation has not been identified yet. On the other hand, s54 activity seems to play a role in exponential silencing. In fact the FtsH protease, involved in maintaining s54 activity levels, is required to reach maximum Pu activity (Carmona and de Lorenzo, 1999). This mechanism has been suggested to influence exponential silencing. Finally, analysis of the carbon metabolism in P. putida showed that Entner-Doudoroff pathway enzyme mutants were impaired in carbon catabolite repression (Vela´zquez et al., 2004). Furthermore, recent analysis of a series of P. putida metabolic mutants has revealed that toluene and glucose exert a reciprocal repression of degradation pathways (del Castillo and Ramos, 2007). Glucose-repression of Pu expression requires ptsN, as it had previously been shown (Cases and de Lorenzo, 2005; Ruı´z et al., 2004). Interestingly, toluene mediated repression of glucose catabolism is mediated by the Crc protein, a key player in Pseudomonads catabolite repression (Collier et al., 1996). Crc has also been shown to play a minor role in rich medium Pu repression (Aranda-Olmedo et al., 2005). Although Crc effect on Pu expression

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. Figure 1 Red arrows indicate a direct positive effect on transcription. Green arrows indicate activation of the regulator. Upwards arrows indicate accumulation of a sigma factor.

has not been analyzed in depth, important clues on the unusual mode of action of this interesting protein have been reported for P. putida alkane degradation pathway in (Moreno et al., 2007). This RNA-binding protein contributes to the control of the system master regulator AlkS by inhibit translation of its messenger after binding to the translation initiation region. Connection with carbon source availability is controlled through Crc abundance by a still unknown mechanism.

5

Research Needs and Future Perspectives

After many years of thorough research, TOL plasmid catabolic pathways can be considered a perfect example of transcription regulation in bacteria. Future studies will focus on the subtleties of the system, with the aim to identify the minute processes involved in the key steps in transcription activation mechanisms, which will help us to understand transcription regulation in bacteria. This will include the basic mechanisms leading to s54 promoter activation, the contacts established between XylS and the two RNAP at the Pm promoter to initiate transcription, and the identification and analysis of new post-transcriptional regulation mechanisms (Vela´zquez et al., 2005). Understanding the processes underlying global control will definitely be the subject of thorough analysis, especially to determine the role of the C/N PTS system, the final target of this phosphor-relay and the possible additional elements involved. A general issue barely addressed to date is how highly hydrophobic signal compounds are spread in aqueous systems, i.e., how toluene finds XylR in the cell. Technical difficulties have

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hampered effective approaches to this problem, but future technologies will allow new strategies aimed at understanding the process to be designed.

Acknowledgments This work was supported by grants MMA, 116/2004/3, 1.2-117/2005/2-B, 308-2006/2-1.2 from the Ministerio de Medio Ambiente and BMC2001–0515 from the Ministerio de Ciencia y Tecnologı´a. Authors wish to thank M. T. Gallegos and J. L. Ramos for critical reading of the MS.

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Ramos JL, Stolz A, Reineke W, Timmis KN (1986) Altered effector specificities in regulators of gene expression: TOL plasmid xylS mutants and their use to engineer expansion of the range of aromatics degraded by bacteria. Proc Natl Acad Sci USA 83: 8467–8471. Rappas M, Schumacher J, Beuron F, Niwa H, Bordes P, Wigneshweraraj S, Keetch CA, Robinson CV, Buck M, Zhang X (2005) Structural insights into the activity of enhancer-binding proteins. Science 307: 1972–1975. Rhee S, Martin RG, Rosner JL, Davies DR (1998) A novel DNA-binding motif in MarA: the first structure for an AraC family transcriptional activator. Proc Natl Acad Sci USA 95: 10413–10418. Ruı´z R, Aranda-Olmedo MI, Domı´nguez-Cuevas P, Ramos-Gonza´lez MI, Marque´s S (2004) Transcriptional regulation of the toluene catabolic pathways. In Pseudomonas. JL Ramos (ed.). London: Kluwer, pp. 509–537. Ruı´z R, Marque´s S, Ramos JL (2003) Leucines 193 and 194 at the N-terminal domain of the XylS protein, the positive transcriptional regulator of the TOL meta-cleavage pathway, are involved in dimerization. J Bacteriol 185: 3036–3041. Ruı´z R, Ramos JL (2002) Residues 137 and 153 at the N terminus of the XylS protein influence the effector profile of this transcriptional regulator and the sigma factor used by RNA polymerase to stimulate transcription from its cognate promoter. J Biol Chem 277: 7282–7286. Salto R, Delgado A, Micha´n C, Marque´s S, Ramos JL (1998) Modulation of the function of the signal receptor domain of XylR, a member of a family of prokaryotic enhancer-like positive regulators. J Bacteriol 180: 600–604. Sallai L, Tucker PA (2005) Crystal structure of the central and C-terminal domain of the sigma(54)-activator ZraR. J Struct Biol 151: 160–170. Schumacher J, Joly N, Rappas M, Zhang X, Buck M (2006) Structures and organisation of AAA+ enhancer binding proteins in transcriptional activation. J Struct Biol 156: 190–199. Shingler V, Pavel H (1995) Direct regulation of the ATPase activity of the transcriptional activator DmpR by aromatic compounds. Mol Microbiol 17: 505–513. Studholme DJ, Dixon R (2003) Domain architectures of sigma54-dependent transcriptional activators. J Bacteriol 185: 1757–1767. Suh SJ, Runyen-Janecky LJ, Maleniak TC, Hager P, MacGregor CH, Zielinski-Mozny NA, Phibbs PV Jr., West SE (2002) Effect of vfr mutation on global gene expression and catabolite repression control of Pseudomonas aeruginosa. Microbiology 148: 1561–1569.

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Sze CC, Shingler V (1999) The alarmone (p)ppGpp mediates physiological-responsive control at the sigma 54-dependent Po promoter. Mol Microbiol 31: 1217–1228. Tobes R, Ramos JL (2002) AraC-XylS database: a family of positive transcriptional regulators in bacteria. Nucleic Acids Res 30: 318–321. Valls M, Buckle M, de Lorenzo V (2002) In vivo UV laser footprinting of the Pseudomonas putida s54 Pu promoter reveals that integration host factor couples transcriptional activity to growth phase. J Biol Chem 277: 2169–2175. Vela´zquez F, di Bartolo I, de Lorenzo V (2004) Genetic evidence that catabolites of the Entner-Doudoroff pathway signal C source repression of the sigma54 Pu promoter of Pseudomonas putida. J Bacteriol 186: 8267–8275. Vela´zquez F, Ferna´ndez S, de Lorenzo V (2006) The upstream-activating sequences of the sigma54

promoter Pu of Pseudomonas putida filter transcription readthrough from upstream genes. J Biol Chem 281: 11940–11948. Vela´zquez F, Parro V, de Lorenzo V (2005) Inferring the genetic network of m-xylene metabolism through expression profiling of the xyl genes of Pseudomonas putida mt-2. Mol Microbiol 57: 1557–1569. Wigneshweraraj S, Bose D, Burrows PC, Joly N, Schumacher J, Rappas M, Pape T, Zhang X, Stockley P, Severinov K, Buck M (2008) Modus operandi of the bacterial RNA polymerase containing the sigma54 promoter-specificity factor. Mol Microbiol 68: 538–546. Williams PA, Murray K (1974) Metabolism of benzoate and the methylbenzoates by Pseudomonas putida (arvilla) mt 2: evidence for the existence of a TOL plasmid. J Bacteriol 120: 416–423.

23 Genetic Features and Regulation of n-Alkane Metabolism F. Rojo Centro Nacional de Biotecnologı´a, CSIC, Cantoblanco, Madrid, Spain [email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1142

2 Regulation of the Expression of Alkane Degradation Genes . . . . . . . . . . . . . . . . . . . . . 1143 2.1 Specific Regulators: Are There Common Characteristics? . . . . . . . . . . . . . . . . . . . . . . . . . . 1143 2.2 Regulation of Pseudomonas butanovora Genes Coding for Butane Monooxygenase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1144 2.3 Regulation of Pseudomonas putida GPo1 Alkane Degradation Pathway . . . . . . . . . . . . 1145 2.4 Why Alkanes are not Preferred Growth Substrates? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1148 2.5 Differential Regulation of Multiple Alkane Hydroxylases . . . . . . . . . . . . . . . . . . . . . . . . . . 1148 2.6 Expression of the Different Components of Alkane Hydroxylases is not Always Coordinated . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1150 3

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1151

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_79, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Expression of the bacterial genes involved in the assimilation of alkanes is frequently tightly regulated. Regulators responding to the presence of alkanes assure that the alkane-degradation genes are induced only when these hydrocarbons are available to the cell. In those microorganisms containing several sets of alkane degradation genes, each one being active on a particular kind of alkanes, regulators should assure a proper differential induction of each set of genes. In addition, the expression of alkane degradation pathways is often downmodulated by a complex global regulation control that assures that the genes are expressed only under the appropriate physiological conditions, or when no other preferred compound are present. The final picture is therefore rather complex and shows that alkane degradation pathways can be highly integrated within cell physiology, suggesting that they can play an important role for survival in many environments.

1

Introduction

The ability to assimilate alkanes is widespread among microorganisms. This may be due, at least in part, to the fact that alkanes are highly reduced molecules with a high energy and carbon content, and therefore can be good carbon and energy sources for those microorganisms able to metabolize them. In addition, besides being major constituents of crude oil, alkanes are produced in low amounts by many living organisms such as plants, green algae, bacteria and animals. This leads to alkanes being present in low quantities in most soils and waters, probably maintaining low but constant concentrations. The biochemistry of alkanedegradation pathways has been studied in detail for several microorganisms (See > Chapter 3, Vol. 2, Part 2), and the genes involved have been identified in many cases. However, much less is known about how the expression of these genes is regulated. Where analyzed, and with some exceptions, expression of the genes involved in the assimilation of alkanes has been found to be tightly regulated. A specific regulator assures that the pathway genes are expressed only in the presence of alkanes. In addition, superimposed to this specific regulation there is frequently a complex control, mediated by global regulators, which assures that the pathways are induced only under appropriate physiological or environmental conditions. For example, when in addition to alkanes cells are faced to other potentially assimilable compounds, global control networks can coordinate the induction of the catabolic pathways for each compound to assure a hierarchical assimilation of the individual carbon sources, a process termed catabolite repression. This is particularly important since these hydrocarbons are normally not preferred growth substrates. Many microorganisms can have up to five or more alkane degradation systems, each one being active on alkanes of a certain chain-length or being expressed under specific physiological conditions. In these cases, the regulatory mechanisms should assure an appropriate differential expression of each set of enzymes. It is clear, therefore, that the regulatory processes that control the expression of alkane degradation genes can be very complex. This chapter summarizes what is currently known about these regulatory mechanisms, which is rather limited. Available information, however, shows that these mechanisms can be intricate and are frequently intimately linked to several aspects of cell physiology.

Genetic Features and Regulation of n-Alkane Metabolism

23

2

Regulation of the Expression of Alkane Degradation Genes

2.1

Specific Regulators: Are There Common Characteristics?

Identification of the specific regulators that induce alkane degradation genes in response to the presence of alkanes has been hampered by the frequent lack of clustering of alkane degradation genes. The few regulators characterized belong to different families, namely to the LuxR/MalT, the AraC/XylS, or to other non-related families or regulators (see > Table 1). Proteins of the LuxR family have a short conserved helix-turn-helix DNA binding domain in their C-terminus, and a variable N-terminal domain that receives an activating signal from an effector or from a sensor protein (Fuqua et al., 1994). Some of these regulators are grouped within the MalT subfamily because, like MalT, have an unusually long N-terminal domain that includes an ATP binding site. At least in the case of MalT, transcriptional activation requires ATP (but not ATP hydrolysis) and an effector, which triggers a multimerization of the protein (Schreiber and Richet, 1999). Regulators of the AraC/XylS family are widespread and have two domains, one of which is conserved and contains determinants for DNA binding and

. Table 1 Transcriptional regulators known or presumed to control the expression of alkane degradation pathways Bacterium

Gene

P. putida GPo1

alkS

Family LuxR/MalT

Effector C6–C10 n-alkanes

Evidence Direct

Reference Panke et al. (1999), Sticher et al. (1997)

P. putida P1

alkS

LuxR/MalT

Not tested

Similarity van Beilen et al. (2001)

A. borkumensis SK2

alkS

LuxR/MalT

Not tested

Similarity Schneiker et al. (2006)

A. borkumensis SK2

gntR

GntR

Not tested

No

Schneiker et al. (2006)

A. borkumensis SK2

araC

AraC/XylS

Not tested

No

Schneiker et al. (2006)

A. borkumensis AP1

alkS

LuxR/MalT

Not tested

Similarity van Beilen et al. (2004)

P. butanovora

bmoR s54Dependent

C2–C8 n-alkanols

Direct

Kurth et al. (2008)

P. aeruginosa RR1 gntR

GntR

C10-C20 n-alkanes

Indirect

Marı´n et al. (2003)

Acinetobacter sp. ADP1

alkR

AraC/XylS

C7-C18 n-alkanes

Direct

Ratajczak et al. (1998)

Acinetobacter sp. M1

alkRa AraC/XylS

>C22 n-alkanes

Indirect

Tani et al. (2001)

Acinetobacter sp. M1

alkRb OruR

C16–C22 n-alkanes

Indirect

Tani et al. (2001)

Global regulators are not included

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Genetic Features and Regulation of n-Alkane Metabolism

transcription activation, while the non-conserved domain is critical for signal recognition in members of the family activated by binding of an effector (Gallegos et al., 1997). For some of these regulators there is evidence supporting that n-alkanes or n-alkanols act as effectors. The water solubility of alkanes having more than eight carbon atoms is below the micromolar range and most likely accumulate into the cytoplasmic membrane. Transcriptional regulators are normally cytoplasmic proteins, since they should interact with DNA. The question arises as to how these regulators interact with the alkanes. The Alcanivorax borkumensis AlkS transcriptional regulator is believed to activate expression of the gene coding for the AlkB1 alkane hydroxylase and of downstream genes in response to alkanes (Schneiker et al., 2006; van Beilen et al., 2004). In a proteomic study this regulator appeared associated to the membrane fraction, rather than to the cytoplasmic fraction (Sabirova et al., 2006). Although AlkS does not show the characteristics expected for a membrane protein, it may have affinity for the inner side of the cytoplasmic membrane, where it has an easy access to the alkanes acting as effectors. Once bound to the alkane, AlkS should move and find its binding site on the DNA. The possible changes that the alkane effector can induce on AlkS have not been reported. The membrane affinity of other alkane-responsive regulators has not been analyzed either.

2.2

Regulation of Pseudomonas butanovora Genes Coding for Butane Monooxygenase

Pseudomonas butanovora can oxidize C2–C8 n-alkanes into the corresponding alcohols utilizing an alkane monooxygenase termed butane monooxygenase (BMO); the alcohols generated are assimilated via oxidation to aldehydes and fatty acids (Arp, 1999). BMO is a multimeric protein formed by the products of the bmoXYBZDC operon (Sluis et al., 2002; see > Fig. 1). Expression of the genes coding for BMO is activated by BmoR, a s54-dependent transcriptional regulator that recognizes as effectors the alcohols and aldehydes derived from the C2–C8 n-alkanes that are substrates of BMO, but not the alkanes themselves (Kurth et al., 2008). Activation by BMO products avoids induction by compounds that are not substrates of BMO, but has the disadvantage that requires a constitutive basal expression of the BMO genes to assure the oxidation of the first molecules of substrate alkanes when these become available to the cell. Regulation is complicated by several factors. Propionate, the final product of propane oxidation, acts as a potent repressor of BMO operon transcription (Doughty et al., 2006) and as a direct inhibitor of BMO activity (Doughty et al., 2007), an effect that persists until propionate catabolism is induced. This is a case of ‘‘product repression.’’ Propionate catabolism is inactive during growth on butane, but is activated by the presence of propionate or upon growth on propane or pentane. Regulation of the metabolism of fatty acids in P. butanovora is still unknown. It has been speculated that a FadR-like fatty acid-responsive regulator may coordinate the expression of the genes coding for BMO with those coding for the degradation of fatty acids. However, the molecular details remain to be elucidated. Repression of alkane hydroxylase genes by the fatty acids generated through oxidation of the corresponding alkanes has been observed in other bacterial strains as well (Marı´n et al., 2001, 2003). Expression of the BMO operon is also induced in response to carbon starvation by an unknown mechanism that is independent of BmoR (Doughty et al., 2007). Finally, it is interesting that expression of the bmoXYBZDC genes generates a BMO that is inactive and that requires activation by the chaperonin-like protein BmoG (Kurth et al., 2008).

Genetic Features and Regulation of n-Alkane Metabolism

23

. Figure 1 Regulation of the genes coding for butane monooxygenase (BMO) in P. butanovora.

2.3

Regulation of Pseudomonas putida GPo1 Alkane Degradation Pathway

Pseudomonas putida GPo1 harbors a plasmid, named OCT, that encodes all genes required for the assimilation of C3–C13 alkanes (Johnson and Hyman, 2006; van Beilen et al., 1994, 2005). The genes of this pathway are grouped in two clusters, alkBFGHJKL and alkST (van Beilen et al., 1994, 2001; see > Fig. 2). The first enzyme of the pathway is an alkane hydroxylase formed by three components, an integral membrane monooxygenase (AlkB) and two soluble proteins, rubredoxin (AlkG) and rubredoxin reductase (AlkT), which transfer electrons from NADH to the monooxygenase. The alkBFGHJKL operon codes for two of the three components of the alkane hydroxylase, and for enzymes involved in further metabolic steps. The alkT gene, encoding the third component of the alkane hydroxylase, is located in the second cluster downstream of alkS, which codes for the transcriptional regulator of the pathway. The alkBFGHJKL operon is transcribed from a promoter, named PalkB, whose expression requires the transcriptional activator AlkS and the presence of alkanes, which act as effectors for AlkS (Kok et al., 1989; Panke et al., 1999). In the absence of alkanes, the alkST genes are expressed at low levels from promoter PalkS1, levels that are negatively regulated by binding of AlkS to the PalkS1 – 10 promoter element (Canosa et al., 1999, 2000). When alkanes become available, AlkS binds to (and represses) PalkS1 more efficiently and, from this site, activates promoter PalkS2, located 38 nt downstream from PalkS1 and which provides high expression of the alkST genes (Canosa et al., 2000). Therefore, the pathway is controlled by a positive feedback mechanism governed by AlkS.

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Genetic Features and Regulation of n-Alkane Metabolism

. Figure 2 Regulation of the genes coding for the alkane degradation pathway of P. putida GPo1.

Promoter PalkS1 is recognized by a form RNA polymerase bound to the alternative sigma factor sS (Canosa et al., 1999). This sigma factor is preferentially expressed under diverse stress conditions, such as during starvation or in stationary phase (Ramos-Gonza´lez and Molin, 1998; Venturi, 2003; Yuste et al., 2006). Therefore, promoter PalkS1 facilitates expression of alkS when cells have neither alkanes nor other carbon sources available, providing enough AlkS levels to allow for a fast response if alkanes eventually enter the cell. On the other hand, the AlkS-dependent promoters PalkS2 and PalkB are probably recognized by s70-RNA polymerase. An AlkS-dependent reporter system based on a PalkB-luxAB fusion showed that C5–C10 alkanes are efficient effectors for the AlkS regulator, while shorter or larger alkanes are not (Sticher et al., 1997). However, P. putida GPo1 can also grow on C3–C4 and C11–C13 alkanes, although significantly more slowly and with a considerable lag time (Johnson and Hyman, 2006; van Beilen et al., 2005), probably because these alkanes are poor inducers of the pathway genes. When using an efficient effector, such as octane, AlkS can respond to alkane concentrations as low as 25 nM (Sticher et al., 1997). It is worth noting that the non-metabolizable analogue dicyclopropylketone can also act as effector of AlkS and as inducer of the pathway genes (Grund et al., 1975). Activation of promoters PalkB and PalkS2 by AlkS and alkanes is negatively modulated by a dominant global control when cells grow in a complete medium, or in a minimal salts medium containing other alternative carbon sources such as amino acids, succinate or lactate (Canosa et al., 2000; Staijen et al., 1999; Yuste et al., 1998). Compounds such as citrate, pyruvate or glycerol, which are also metabolized, do not exert this inhibitory effect. Repression is particularly strong during exponential growth in a complete medium (about 70-fold inhibition as measured with a PalkB-lacZ transcriptional fusion), but rapidly fades away when cells enter into stationary phase. The repression exerted by succinate or lactate in a minimal salts medium is significantly milder, in the range of 4- to 5-fold (Yuste et al., 1998). Repression is not related to the growth rate that each carbon source allows. There is no clear correlation between growth rate and the extent of inhibition exerted by each carbon source (Yuste et al., 1998). In addition, the use of chemostats showed a clear repression in cultures in which the growth rate was controlled by limiting the source of nitrogen but that contained succinate in excess as the carbon source. However, repression was not observed when the same growth rate was obtained by limiting the availability of succinate, keeping all other nutrients in excess (Dinamarca et al., 2003). Therefore, it is the presence of an excess of succinate what

Genetic Features and Regulation of n-Alkane Metabolism

23

inhibits expression of the alkane degradation pathway. The repressive signals probably arise from the levels of key internal metabolites or compounds, which vary according to the carbon source being used and on its concentration in the medium. When cells grow in a complete medium, the negative control depends on the additive effects of two global regulation networks. One of them relies on the global regulatory protein Crc (Yuste and Rojo, 2001), while the other one receives information from the cytochrome o ubiquinol oxidase (Cyo), a component of the electron transport chain (Dinamarca et al., 2002, 2003). However, when cells grow in a minimal salts medium containing succinate as the carbon source, the effect of Crc is very small and most of the global control inhibiting the induction of the alkane degradation genes derives from the Cyo terminal oxidase (Dinamarca et al., 2003; Yuste and Rojo, 2001). At least in a complete medium, the inhibition process generates a strong decrease in the levels of the AlkS transcriptional activator, an unstable protein present in the cell in limiting amounts even under inducing conditions (Yuste and Rojo, 2001). By keeping AlkS levels below those required for maximal induction of the pathway, expression of the alkST and alkBFGHJKL operons can be down-modulated in a simple and coordinated way (see > Fig. 1). Since AlkS activates expression of its own gene, its levels could be modulated by limiting alkS transcription from promoter PalkS2, by inhibiting translation of alkS mRNA, or by controlling alkS mRNA stability. All these alternatives would lead to the same final result, this is, to decreased levels of the AlkS protein. Crc seems to have little influence on alkS mRNA stability (Yuste and Rojo, 2001). A detailed analysis showed that Crc is an RNA-binding protein that interacts with the 50 -end of alkS mRNA, inhibiting translation (Moreno et al., 2007). Decreasing translation of alkS indirectly leads to reduced alkS transcription, given that AlkS activates the expression of its own gene from promoter PalkS2. Crc inhibits as well the expression of many other catabolic pathways for several non-preferred compounds in Pseudomonads (Aranda-Olmedo et al., 2005; Hester et al., 2000a, b; MacGregor et al., 1996; Morales et al., 2004). In at least one other example it has been demonstrated that Crc acts by inhibiting translation of the mRNA of the target gene (Moreno and Rojo, 2008). The levels and activity of Crc vary depending on growth conditions (Ruiz-Manzano et al., 2005), but signal that triggers Crc activity is not known. The mechanism through which Cyo influences the expression of the alkane degradation pathway is still unclear. Cyo is one of the five terminal oxidases characterized in P. putida. The differential expression of these terminal oxidases is carefully regulated in a coordinated fashion to optimize energy production under the prevailing environmental conditions (Ugidos et al., 2008; Williams et al., 2007). Inactivation of the Cyo terminal oxidase partially relieves the repression exerted on the alkane degradation pathway under several conditions, while inactivation of any of the other four terminal oxidases does not (Dinamarca et al., 2002; Morales et al., 2006). The absence of Cyo affects the expression of many other genes, so that it has been proposed to be a component of a global regulation network that transmits information on the activity of the electron transport chain to coordinate respiration and carbon metabolism (Morales et al., 2006; Petruschka et al., 2001). Expression of the genes coding for the Cyo terminal oxidase varies strongly according to oxygen levels and the carbon source being used and there is a clear correlation between Cyo levels and the extent of repression of the alkane degradation pathway (Dinamarca et al., 2003). However, how Cyo transmits the signal that eventually triggers a process of regulation of gene expression, which are the components that directly exert the regulatory effect, and which is the precise mechanism, are questions that remain unsolved.

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Genetic Features and Regulation of n-Alkane Metabolism

Why Alkanes are not Preferred Growth Substrates?

With a few exceptions, hydrocarbons are not preferred growth substrates for bacterial strains having a versatile metabolism. In particular, the expression of alkane degradation pathways is inhibited by the presence of many other carbon sources in several bacterial strains of P. putida, P. aeruginosa, B. cepacia, P. butanovora or Acinetobacter sp. (Doughty et al., 2006; Marı´n et al., 2001, 2003; Ratajczak et al., 1998; Staijen et al., 1999; Yuste et al., 1998). The preferential use of a carbon source over other potential substrates when these are all present in the growth medium is a global regulation phenomenon generally termed as catabolite repression control (Magasanik, 1970). However, the preferred compounds vary in distinct bacterial species and the mechanisms used to modulate the expression of catabolic pathways, where characterized, frequently differ as well (Rojo and Dinamarca, 2004). It is likely, therefore, that this process arises from a number of global regulatory mechanisms that are directed to optimize carbon metabolism and energy generation in response to different signals. Which these signals are is not clear, but may be related to the concentrations of key metabolites or molecules in the cell that in turn depend on the efficiency of the different catabolic pathways in terms of energy gain. Alkanes are generally considered as non-toxic compounds. However, there are several reports indicating that the use of alkanes as a carbon source can have deleterious consequences to cell physiology that force an adaptation process. Growth of P. putida GPo1 using octane as the carbon source modifies the fatty acid composition of the cytoplasmic membrane as well as cell morphology and viability (Chen et al., 1995). Surprisingly, this was traced in part to the presence of the AlkB alkane hydroxylase in the membrane. Induction of the alkB gene by the gratuitous inducer dicyclopropylketone, rather than octane, was detrimental to cell physiology. Growth of cells in a complete medium with dicyclopropylketone for some generations led to the isolation of mutant derivatives in which alkB could no longer be induced, an effect that was not observed if dicyclopropylketone was omitted (Chen et al., 1996). Apparently, the AlkB alkane hydroxylase and the AlkJ alcohol dehydrogenase are produced to very high amounts in cells growing at the expense of octane and this affects the cell membrane. The n-octanol generated by AlkB from octane also has an effect on cell membrane composition, leading to an increase in trans unsaturated fatty acids (Chen et al., 1995). These results are interesting in the context of the inhibition that fatty alcohols and fatty acids generate on expression of alkane degradation genes in several bacterial strains (Doughty et al., 2006; Marı´n et al., 2001, 2003; Ratajczak et al., 1998), suggesting that it represents a ‘‘product repression’’ effect to coordinate the generation of these compounds with their further metabolism, thereby avoiding their deleterious accumulation in the cell membrane.

2.5

Differential Regulation of Multiple Alkane Hydroxylases

Several bacterial strains contain more than one alkane hydroxylase and in some cases it has been shown that the corresponding genes are differentially regulated. Acinetobacter sp. strain M-1 contains two AlkB-related alkane hydroxylases named AlkMa and AlkMb. It is not clear which are the enzymatic differences between the two proteins, but they are differentially regulated depending on the alkane used as carbon source. Expression of AlkMa, which is controlled by the AlkRa regulator, is induced by alkanes having a very long chain length (>C22), while that of AlkMb is induced by AlkRb in the presence of C16–C22 alkanes (Tani et al., 2001).

Genetic Features and Regulation of n-Alkane Metabolism

23

P. aeruginosa strains RR1 and PAO1 contain two alkane hydroxylases, AlkB1 and AlkB2. At least in strain PAO1, the substrate range of the two enzymes overlaps significantly, since AlkB1 oxidizes C16–C24 n-alkanes while AlkB2 is active on C12–C20 n-alkanes. The regulation of the genes encoding for AlkB1 and AlkB2 has been studied in strain RR1 (Marı´n et al., 2003). The expression of both genes is induced by C10–C22 alkanes, although transcription of alkB1 is almost twice as efficient as that of alkB2. The alkB2 gene is induced preferentially during the early exponential phase of growth, while alkB1 is induced in the late-exponential phase of growth. The expression of both genes declines in stationary phase. The regulators responsible for this differential regulation have not been characterized, although a gene coding for a GntRlike regulator maps immediately upstream of alkB2. Some bacterial strains have four or more alkane oxidation systems. An interesting example is that of A. borkumensis, which is highly specialized in metabolizing alkanes and has two AlkB-like alkane hydroxylases and three genes coding for cytochromes P450 believed to be involved in alkane oxidation (Hara et al., 2004; Schneiker et al., 2006; van Beilen et al., 2004). The organization of these genes is depicted in > Fig. 3. The substrate range of these alkane hydroxylation systems partially overlaps. AlkB1 oxidizes C5–C12 n-alkanes, while AlkB2 is active on C8–C16 n-alkanes (van Beilen et al., 2004). They probably share the auxiliary proteins rubredoxin and rubredoxin reductase, which are encoded by genes (rubA and rubB) that map separately from alkB1 and alkB2. In A. borkumensis strain AP1, expression of the alkB1 and alkB2 genes is very low when cells grow using pyruvate as the carbon source, but is strongly induced when C10–C16 alkanes are metabolized (van Beilen et al., 2004). Expression decreases considerably upon entry into stationary phase, a behavior that has been observed as well in the A. borkumensis strain SK2 (Schneiker et al., 2006). A proteomic analysis showed that the expression of genes alkB1, alkG, alkH and alkJ, all of which are involved in alkane oxidation and presumably form an operon, is up-regulated in the presence of hexadecane (Sabirova et al., 2006). Upstream from alkB1 there is a gene showing similarity to P. putida GPo1 alkS that is predicted to be an alkane-responsive transcriptional activator. Transcription of A. borkumensis AP1 alkS seems to be constitutive (van Beilen et al., 2004), contrary to what was observed for

. Figure 3 Genes involved in the oxidation of n-alkanes in A. borkumensis SK2. See text for details.

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Genetic Features and Regulation of n-Alkane Metabolism

P. putida GPo1 alkS. The proteomic analysis performed in strain SK2 detected higher AlkS levels in hexadecane-grown cells than in pyruvate-grown cells (Sabirova et al., 2006), although it is unknown whether this is due to increased transcription of alkS, to a higher stability of the AlkS protein in the presence of alkanes, or to other reasons. The promoter for the A. borkumensis alkB1 gene contains a good alkS-binding site immediately upstream of the 35 promoter element, as it occurs in P. putida GPo1 alkB, but a similar binding site is not evident at the promoter for alkB2 (van Beilen et al., 2004). P. putida GPo1 AlkS could substitute for A. borkumensis AP1 at activating expression of the alkB1 gene, but the same was not true for the alkB2 gene. Therefore, available evidence suggests that A. borkumensis AlkS activates expression of the alkB1 alkane hydroxylase in response to alkanes, but is unlikely to regulate expression of alkB2, which is nevertheless inducible by alkanes. Interestingly, a gene coding for a transcriptional regulator of the GntR family is located just upstream of alkB2, although its role in alkB2 expression has not been reported. A. borkumensis has three genes encoding proteins with significant sequence similarity to cytochromes P450 of the CYP153 family (Schneiker et al., 2006); cytochromes of this family are involved in alkane oxidation in other microorganisms (Funhoff et al., 2006; Maier et al., 2001; van Beilen et al., 2006). Cytochrome P450-1 maps adjacent to genes coding for a ferredoxin (fdx), for the AlkJ2 alcohol dehydrogenase and for a putative oxidoreductase (> Fig. 3). Cytochrome P450-2 is identical to P450-1, and highly homologous to P450-3. Proteomic profiling analyses revealed that P450-1 and/or P450-2, which cannot be differentiated with this technique, are expressed in cells grown with either pyruvate or hexadecane as the carbon source, although expression was higher in alkane-grown cells (Sabirova et al., 2006). Since AlkJ2 protein was also up-regulated by hexadecane and probably forms an operon with P450-1, it is likely that expression of P450-1 is induced by hexadecane but that of P450-2 and P450-3 is not. A gene coding for a transcriptional regulator of the AraC family maps close to P450-1, but its role in regulating the P450-1 cluster has not been reported. Although not investigated in detail, the different alkane hydroxylating systems present in A. borkumensis are likely to have different properties in terms of substrate specificity, turnover rate, etc. In addition to the AlkB-like and P450 genes mentioned above, A. borkumensis seems to have other uncharacterized genes involved in oxidation of branched alkanes and phytane (Schneiker et al., 2006). Finally, a gene similar to Acinetobacter sp. DSM17874 almA, which oxidizes alkanes of very long chain-length, has been predicted in A. borkumensis SK2 (ThroneHolst et al., 2007). Expression of all these alkane oxidation genes should be differentially induced according to the substrate present under each circumstance, although the regulators involved and/or the signals to which they respond remain unknown.

2.6

Expression of the Different Components of Alkane Hydroxylases is not Always Coordinated

As mentioned above, alkane hydroxylases of the AlkB family are formed by an integralmembrane monooxygenase (commonly named as ‘‘alkane hydroxylase’’) and two soluble auxiliary proteins, rubredoxin and rubredoxin reductase, which transfer electrons from NADH to the monooxygenase. In some cases, such as in P. putida GPo1, expression of the genes coding for the rubredoxin and rubredoxin reductase is coordinated with that of the AlkB alkane monooxygenase; the three genes are controlled by the AlkS transcriptional regulator and are up-regulated by alkanes (reviewed in van Beilen et al., 2001). However, there are

Genetic Features and Regulation of n-Alkane Metabolism

23

several bacterial strains in which this is not the case. For example, rubredoxin and rubredoxin reductase are constitutively expressed in Acinetobacter sp. strains M-1 (Tani et al., 2001) and ADP1 (Geissdorfer et al., 1999), as well as in P. aeruginosa RR1 and PAO1 (Marı´n et al., 2003), while expression of the genes coding for the corresponding AlkB-like alkane monooxygenases is induced by alkanes. It is unclear whether in these later cases the rubredoxin and rubredoxin reductase are shared with other monooxygenases.

3

Research Needs

While there is no doubt that the expression of most alkane degradation genes is regulated, how regulation is accomplished is unclear in most cases. The specific regulators responsible for the induction of these pathways have been identified in only very few examples. Their characterization is important, especially in those microorganisms that have several alkane hydroxylation systems, each one with particular albeit sometimes overlapping specificities, and that should be differentially regulated. A particularly important case is that of hydrocarbonoclastic bacteria such as A. borkumensis, because of their specialization in alkane metabolism and because they contain several independent alkane hydroxylation systems. An intriguing question is why the expression of the monooxygenase component of alkane hydroxylases is invariably regulated, while that of the auxiliary proteins rubredoxin and rubredoxin reductase is constitutive in many bacteria. Perhaps they serve other functions in addition to transferring electrons to alkane monooxygenases, but this has not been clarified. It has been important to realize that the expression of the alkane degradation pathways is frequently coordinated with other aspects of cell metabolism. Most efforts have been directed to elucidate the mechanisms responsible for the catabolite repression control, but there are other global regulation phenomena that modulate the expression of alkane degradation genes. The picture is far from clear and requires much more efforts in different microorganisms, because the molecular mechanisms will likely be different in each case. Finally, there are indications that the expression of alkane degradation pathways is coordinated with the induction of fatty acid metabolism and with generation of storage polymers, but the details on this are again missing in most cases.

Acknowledgments The critical comments of R. Moreno are gratefully acknowledged. Work on author’s lab was funded by grants BFU2006-00767/BMC and CSD2007-00005.

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Canosa I, Sa´nchez-Romero JM, Yuste L, Rojo F (2000) A positive feedback mechanism controls expression of AlkS, the transcriptional regulator of the Pseudomonas oleovorans alkane degradation pathway. Mol Microbiol 35: 791–799. Canosa I, Yuste L, Rojo F (1999) Role of the alternative sigma factor sigmaS in expression of the AlkS

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regulator of the Pseudomonas oleovorans alkane degradation pathway. J Bacteriol 181: 1748–1754. Chen Q, Janssen DB, Witholt B (1995) Growth on octane alters the membrane lipid fatty acids of Pseudomonas oleovorans due to the induction of alkB and synthesis of octanol. J Bacteriol 177: 6894–6901. Chen Q, Janssen DB, Witholt B (1996) Physiological changes and alk gene instability in Pseudomonas oleovorans during induction and expression of alk genes. J Bacteriol 178: 5508–5512. Dinamarca MA, Aranda-Olmedo I, Puyet A, Rojo F (2003) Expression of the Pseudomonas putida OCT plasmid alkane degradation pathway is modulated by two different global control signals: evidence from continuous cultures. J Bacteriol 185: 4772–4778. Dinamarca MA, Ruiz-Manzano A, Rojo F (2002) Inactivation of cytochrome o ubiquinol oxidase relieves catabolic repression of the Pseudomonas putida GPo1 alkane degradation pathway. J Bacteriol 184: 3785–3793. Doughty DM, Halsey KH, Vieville CJ, Sayavedra-Soto LA, Arp DJ, Bottomley PJ (2007) Propionate inactivation of butane monooxygenase activity in ‘‘Pseudomonas butanovora’’: biochemical and physiological implications. Microbiology 153: 3722–3729. Doughty DM, Sayavedra-Soto LA, Arp DJ, Bottomley PJ (2006) Product repression of alkane monooxygenase expression in Pseudomonas butanovora. J Bacteriol 188: 2586–2592. Funhoff EG, Bauer U, Garcia-Rubio I, Witholt B, van Beilen JB (2006) CYP153A6, a soluble P450 oxygenase catalyzing terminal-alkane hydroxylation. J Bacteriol 188: 5220–5227. Fuqua WC, Winans SC, Greenberg EP (1994) Quorum sensing in bacteria: the LuxR-LuxI family of cell density-responsive transcriptional regulators. J Bacteriol 176: 269–275. Gallegos MT, Schleif R, Bairoch A, Hofmann K, Ramos JL (1997) Arac/XylS family of transcriptional regulators. Microbiol Mol Biol Rev 61: 393–410. Geissdorfer W, Kok RG, Ratajczak A, Hellingwerf KJ, Hillen W (1999) The genes rubA and rubB for alkane degradation in Acinetobacter sp. strain ADP1 are in an operon with estB, encoding an esterase, and oxyR. J Bacteriol 181: 4292–4298. Grund A, Shapiro J, Fennewald M, Bacha P, Leahy J, Markbreiter K, Nieder M, Toepfer M (1975) Regulation of alkane oxidation in Pseudomonas putida. J Bacteriol 123: 546–556. Hara A, Baik SH, Syutsubo K, Misawa N, Smits TH, van Beilen JB, Harayama S (2004) Cloning and functional analysis of alkB genes in Alcanivorax borkumensis SK2. Environ Microbiol 6: 191–197.

Hester KL, Lehman J, Najar F, Song L, Roe BA, MacGregor CH, Hager PW, Phibbs PV Jr, Sokatch JR (2000a) Crc is involved in catabolite repression control of the bkd operons of Pseudomonas putida and Pseudomonas aeruginosa. J Bacteriol 182: 1144–1149. Hester KL, Madhusudhan KT, Sokatch JR (2000b) Catabolite repression control by crc in 2xYT medium is mediated by posttranscriptional regulation of bkdR expression in Pseudomonas putida. J Bacteriol 182: 1150–1153. Johnson EL, Hyman MR (2006) Propane and n-butane oxidation by Pseudomonas putida GPo1. Appl Environ Microbiol 72: 950–952. Kok M, Oldenhuis R, van der Linden MP, Raatjes P, Kingma J, van Lelyveld PH, Witholt B (1989) The Pseudomonas oleovorans alkane hydroxylase gene. Sequence and expression. J Biol Chem 264: 5435–5441. Kurth EG, Doughty DM, Bottomley PJ, Arp DJ, Sayavedra-Soto LA (2008) Involvement of BmoR and BmoG in n-alkane metabolism in ‘‘Pseudomonas butanovora.’’ Microbiology 154: 139–147. MacGregor CH, Arora SK, Hager PW, Dail MB, Phibbs PV Jr (1996) The nucleotide sequence of the Pseudomonas aeruginosa pyrE-crc-rph region and the purification of the crc gene product. J Bacteriol 178: 5627–5635. Magasanik B (1970) Glucose effects: inducer exclusion and repression. In The Lactose Operon. J Beckwith (ed.). Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press, pp. 189–220. Maier T, Forster HH, Asperger O, Hahn U (2001) Molecular characterization of the 56-kDa CYP153 from Acinetobacter sp. EB104. Biochem Biophys Res Commun 286: 652–658. Marı´n MM, Smits TH, van Beilen JB, Rojo F (2001) The alkane hydroxylase gene of Burkholderia cepacia RR10 is under catabolite repression control. J Bacteriol 183: 4202–4209. Marı´n MM, Yuste L, Rojo F (2003) Differential expression of the components of the two alkane hydroxylases from Pseudomonas aeruginosa. J Bacteriol 185: 3232–3237. Morales G, Linares JF, Beloso A, Albar JP, Martı´nez JL, Rojo F (2004) The Pseudomonas putida Crc global regulator controls the expression of genes from several chromosomal catabolic pathways for aromatic compounds. J Bacteriol 186: 1337–1344. Morales G, Ugidos A, Rojo F (2006) Inactivation of the Pseudomonas putida cytochrome o ubiquinol oxidase leads to a significant change in the transcriptome and to increased expression of the CIO and cbb3-1 terminal oxidases. Environ Microbiol 8: 1764–1774.

Genetic Features and Regulation of n-Alkane Metabolism Moreno R, Rojo F (2008) The target for the Pseudomonas putida Crc global regulator in the benzoate degradation pathway is the BenR transcriptional regulator. J Bacteriol 190: 1539–1545. Moreno R, Ruiz-Manzano A, Yuste L, Rojo F (2007) The Pseudomonas putida Crc global regulator is an RNA binding protein that inhibits translation of the AlkS transcriptional regulator. Mol Microbiol 64: 665–675. Panke S, Meyer A, Huber CM, Witholt B, Wubbolts MG (1999) An alkane-responsive expression system for the production of fine chemicals. Appl Environ Microbiol 65: 2324–2332. Petruschka L, Burchhardt G, Mu¨ller C, Weihe C, Herrmann H (2001) The cyo operon of Pseudomonas putida is involved in catabolic repression of phenol degradation. Mol Gen Genom 266: 199–206. Ramos-Gonza´lez MI, Molin S (1998) Cloning, sequencing, and phenotypic characterization of the rpoS gene from Pseudomonas putida KT2440. J Bacteriol 180: 3421–3431. Ratajczak A, Geissdorfer W, Hillen W (1998) Expression of alkane hydroxylase from Acinetobacter sp. strain ADP1 is induced by a broad range of n-alkanes and requires the transcriptional activator AlkR. J Bacteriol 180: 5822–5827. Rojo F, Dinamarca MA (2004) Catabolite repression and physiological control. In Pseudomonas, vol. 2. JL Ramos (ed.). New York: Kluwer Academic/Plenum Publishers, pp. 365–387. Ruiz-Manzano A, Yuste L, Rojo F (2005) Levels and activity of the Pseudomonas putida global regulatory protein Crc vary according to growth conditions. J Bacteriol 187: 3678–3686. Sabirova JS, Ferrer M, Regenhardt D, Timmis KN, Golyshin PN (2006) Proteomic insights into metabolic adaptations in Alcanivorax borkumensis induced by alkane utilization. J Bacteriol 188: 3763–3773. Schneiker S, Martins dos Santos VA, Bartels D, Bekel T, Brecht M, Buhrmester J, Chernikova TN, Denaro R, Ferrer M, Gertler C, Goesmann A, Golyshina OV, Kaminski F, Khachane AN, Lang S, Linke B, McHardy AC, Meyer F, Nechitaylo T, Puhler A, Regenhardt D, Rupp O, Sabirova JS, Selbitschka W, Yakimov MM, Timmis KN, Vorholter FJ, Weidner S, Kaiser O, Golyshin PN (2006) Genome sequence of the ubiquitous hydrocarbon-degrading marine bacterium Alcanivorax borkumensis. Nat Biotechnol 24: 997–1004. Schreiber V, Richet E (1999) Self-association of the Escherichia coli transcription activator MalT in the presence of maltotriose and ATP. J Biol Chem 274: 33220–33226. Sluis MK, Sayavedra-Soto LA, Arp DJ (2002) Molecular analysis of the soluble butane monooxygenase from

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‘‘Pseudomonas butanovora.’’ Microbiology 148: 3617–3629. Staijen IE, Marcionelli R, Witholt B (1999) The PalkBFGHJKL promoter is under carbon catabolite repression control in Pseudomonas oleovorans but not in Escherichia coli alk + recombinants. J Bacteriol 181: 1610–1616. Sticher P, Jaspers MC, Stemmler K, Harms H, Zehnder AJ, van der Meer JR (1997) Development and characterization of a whole-cell bioluminescent sensor for bioavailable middle-chain alkanes in contaminated groundwater samples. Appl Environ Microbiol 63: 4053–4060. Tani A, Ishige T, Sakai Y, Kato N (2001) Gene structures and regulation of the alkane hydroxylase complex in Acinetobacter sp. strain M-1. J Bacteriol 183: 1819–1823. Throne-Holst M, Wentzel A, Ellingsen TE, Kotlar HK, Zotchev SB (2007) Identification of novel genes involved in long-chain n-alkane degradation by Acinetobacter sp. strain DSM 17874. Appl Environ Microbiol 73: 3327–3332. Ugidos A, Morales G, Rial E, Williams HD, Rojo F (2008) The coordinate regulation of multiple terminal oxidases by the Pseudomonas putida ANR global regulator. Environ Microbiol 10: 1690–1702 van Beilen JB, Funhoff EG, van Loon A, Just A, Kaysser L, Bouza M, Holtackers R, Rothlisberger M, Li Z, Witholt B (2006) Cytochrome P450 alkane hydroxylases of the CYP153 family are common in alkane-degrading eubacteria lacking integral membrane alkane hydroxylases. Appl Environ Microbiol 72: 59–65. van Beilen JB, Marı´n MM, Smits TH, Ro¨thlisberger M, Franchini AG, Witholt B, Rojo F (2004) Characterization of two alkane hydroxylase genes from the marine hydrocarbonoclastic bacterium Alcanivorax borkumensis. Environ Microbiol 6: 264–273. van Beilen JB, Panke S, Lucchini S, Franchini AG, Ro¨thlisberger M, Witholt B (2001) Analysis of Pseudomonas putida alkane degradation gene clusters and flanking insertion sequences: evolution and regulation of the alk-genes. Microbiology 147: 1621–1630. van Beilen JB, Smits TH, Roos FF, Brunner T, Balada SB, Ro¨thlisberger M, Witholt B (2005) Identification of an amino acid position that determines the substrate range of integral membrane alkane hydroxylases. J Bacteriol 187: 85–91. van Beilen JB, Wubbolts MG, Witholt B (1994) Genetics of alkane oxidation by Pseudomonas oleovorans. Biodegradation 5: 161–174. Venturi V (2003) Control of rpoS transcription in Escherichia coli and Pseudomonas: why so different? Mol Microbiol 49: 1–9.

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Williams HD, Zlosnik JE, Ryall B (2007) Oxygen, cyanide and energy generation in the cystic fibrosis pathogen Pseudomonas aeruginosa. Adv Microb Physiol 52: 1–71. Yuste L, Canosa I, Rojo F (1998) Carbon-sourcedependent expression of the PalkB promoter from the Pseudomonas oleovorans alkane degradation pathway. J Bacteriol 180: 5218–5226. Yuste L, Herva´s AB, Canosa I, Tobes R, Jime´nez JI, Nogales J, Pe´rez-Pe´rez MM, Santero E, Dı´az E,

Ramos JL, de Lorenzo V, Rojo F (2006) Growthphase dependent expression of the Pseudomonas putida KT2440 transcriptional machinery analyzed with a genome-wide DNA microarray. Environ Microbiol 8: 165–177. Yuste L, Rojo F (2001) Role of the crc gene in catabolic repression of the Pseudomonas putida GPo1 alkane degradation pathway. J Bacteriol 183: 6197–6206.

24 Diversity of Naphthalene Biodegradation Systems in Soil Bacteria A. M. Boronin* . I. A. Kosheleva Skryabin Institute of Biochemistry and Physiology of Microorganisms, Russian Academy of Sciences, Pushchino State University, Pushchino, Russia *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1156

2

Naphthalene-Degrading Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1156

3

Naphthalene Catabolic Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1156

4

Naphthalene Catabolic Plasmids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1157

5 5.1

5.3.1 5.3.2 5.3.3 5.3.4 5.3.5

Genetic Systems for Naphthalene Degradation of Pseudomonas spp . . . . . . . . . . 1157 Genetic Systems for Naphthalene Degradation in Archetypal Strains P. putida G7 and NCIB9816-4 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1157 Naphthalene Catabolic Genetic Systems Similar to nah-Genes of the Plasmid NAH7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1158 Naphthalene Catabolic Genes in P. stutzeri AN10 and Pseudomonas sp. ND6 . . . 1158 PAH Catabolic Genes Organization in P. putida BS202 . . . . . . . . . . . . . . . . . . . . . . . . . . 1158 Organization of Naphthalene Catabolic Genes in P. putida AK5 . . . . . . . . . . . . . . . . . 1159 Naphthalene Catabolic Genetic Systems Differ from Archetype Plasmid NAH7 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1160 PAH Catabolic Genes of Comamonas testosteroni Strains . . . . . . . . . . . . . . . . . . . . . . . . . 1160 Naphthalene Catabolic Genes of Ralstonia sp. Strain U2 . . . . . . . . . . . . . . . . . . . . . . . . . 1161 Phenanthrene Catabolic Genes of Burkholderia sp. Strain RP007 . . . . . . . . . . . . . . . . 1161 Degradation of PAH by Sphingomonas sp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1161 Naphthalene Catabolic Genes of Rhodococcus sp . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1162

6

Variability of PAH Catabolic Systems in Naturally Occurring Bacteria . . . . . . 1162

7

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1162

5.2 5.2.1 5.2.2 5.2.3 5.3

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_80, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: The environment is constantly being polluted by polycyclic aromatic hydrocarbons (PAH) originated from different sources. Aromatic hydrocarbons are known to be degraded by a variety of microorganisms. Non homologous genetic systems involving in biodegradation of PAH and encoding isofunctional enzymes have been found in different species of bacteria. Metabolic pathways are constantly evolving in the response on the environmental changes.

1

Introduction

Naphthalene and its substituted derivatives are representatives of PAH commonly identified in environmental samples. Certain PAH are strong human carcinogens leading to widespread interest in the microbial metabolism of these compounds. Degradation of aromatic hydrocarbons by microorganisms is one of the major processes that results in the decontamination of polluted environment. Oxidation of naphthalene is the best example demonstrating how bacteria can degrade aromatic compound as carbon and energy source. Naphthalene has been often used as a model PAH due to high speed of utilization by microorganisms compared to other PAH and the relatively simple structure of the intermediates in the catabolic pathways. Genetic control of the pathways involved in naphthalene degradation has been studied in detail for the Gram-negative Pseudomonas and related species and for some Gram-positive microorganisms.

2

Naphthalene-Degrading Microorganisms

A large number of different strains of algae, fungi, and bacteria are known to degrade PAH. However, PAH degradation capabilities appeared to be associated with members of certain taxa, independent of the origin of the soils from which bacteria were isolated. The pathways for PAH biodegradation have been reported in bacteria from different genera such as Pseudomonas, Sphingomonas, Burkholderia, Mycobacterium, Corynebacterium, Aeromonas, Rhodococcus, and Bacillus. Naphthalene-degrading microorganisms are widely distributed in nature and are easy to isolate from different coal tar and oil contaminated soils. Among indicated above taxonomic groups a high proportion of the PAH-degrading isolates belong to Pseudomonas, Sphingomonas, and Burkholderia strains.

3

Naphthalene Catabolic Pathways

All Pseudomonas strains investigated capable of aerobic naphthalene degradation oxidize naphthalene through salicylic acid. Salicylate can be oxidized to catechol and then cleaved via the meta-pathway (by catechol 2,3-dioxygenase) or via ortho-pathway (by catechol 1,2-dioxygenase) and through gentisic acid. Naphthalene degradation is initiated metabolism of naphthalene by incorporating dioxygen into the aromatic nucleus. The reaction is catalyzed by a multicomponent enzyme system, naphthalene dioxygenase. This system consists of a flavin-containing ferredoxin reductase (NahAa), an iron–sulfur ferredoxin (NahAb), an iron–sulfur dioxygenase, consisting of two large Rieske-type [2Fe–2S]-containing subunits (NahAc) and two small subunits (NahAd). Salicylate is converted into catechol (through the action of salicylate 1-monooxygenase), or into gentisate (through the action of salicylate 5-hydroxylase). Further oxidation of catechol or gentisate results in formation of tricarboxylic acid cycle intermediates.

Diversity of Naphthalene Biodegradation Systems in Soil Bacteria

4

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Naphthalene Catabolic Plasmids

Genetic systems for the catabolism of naphthalene are often found in fluorescent Pseudomonas on large conjugative plasmids. Plasmids carrying naphthalene catabolic genes represent a class of well-characterized degradation plasmids, known collectively as NAH plasmids. All NAH plasmids investigated to date encode a single upper pathway for the conversion of naphthalene to salicylate. Most of NAH plasmids contain the genes for salicylate metabolism via metapathway of catechol oxidation. Naphthalene catabolic plasmids differ in size, restriction patterns and incompatibility groups and revealed a high level of DNA homology among nah genes suggested that these plasmids are related. Most of naphthalene catabolic plasmids belong to incompatibility groups P-9 and P-7 (Boronin, 1992). Phylogenetic analysis of rep and oriV sequences of IncP-9 NAHlike plasmids revealed five subgroups with 7–35% divergence between them (Sevastsyanovich et al., 2008). More often naphthalene catabolic plasmids were identified as members of IncP9d and IncP-9b subgroups. Environmental studies have suggested a wide distribution of IncP9-like replicons in nature and their involvement in natural horizontal gene transfer of the naphthalene degradation trait. Naphthalene degradative plasmids of IncP-7 group are more rarely than IncP-9 and are structurally diverse and do not form any clusters, in distinction from IncP-9 plasmids.

5

Genetic Systems for Naphthalene Degradation of Pseudomonas spp

5.1

Genetic Systems for Naphthalene Degradation in Archetypal Strains P. putida G7 and NCIB9816-4

The nah genes for naphthalene catabolism are organized in the upper pathway operon, which control initial oxidation and subsequent degradation to salicylate (nah1) and the operon for salicylate oxidation and further meta cleavage of catechol (nah2). Both operons in strains NCIB 9816 and PpG7 are carried on conjugative IncP-9 plasmids (pDTG1 and NAH7, respectively). The operon for upper pathway of naphthalene biodegradation consists of genes nahAaAbAcAdBFCQED, encoding degradation of naphthalene to salicylate and the operon for lower pathway includes nahGTHINLOMKJ genes responsible for conversion of salicylate to pyruvate and acetyl coenzyme A. Downstream of the nahJ gene of strain P. putida G7 two additional genes coding for an unknown function (nahX) and chemotaxis toward naphthalene transducer protein (nahY) were found (Habe and Omori, 2003). A methylaccepting chemotaxis protein, NahY, is co transcribed with the degradation genes. Both nah1 and nah2 operons are activated by a trans-acting positive regulator encoded by nahR gene and commonly are induced by naphthalene intermediate – salicylate. The product of nahR gene, NahR is a member of the LysR family of transcriptional activators. The NahR binds to the promoters of nah1 and nah2 operons and activated their transcription after interaction with salicylate (Shell, 1990). NahR proteins from different naphthalene-degrading P. putida strains exhibit a highly conserved helix-turn-helix motif and a putative enhancer-binding region in the N-terminal domain. Gene activation by NahR is consistent with general transcriptional mechanism of class I transcription factors, by protein–protein interactions between alphaRNAP and the NahR (Park et al., 2002).

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Naphthalene Catabolic Genetic Systems Similar to nah-Genes of the Plasmid NAH7

Nucleotide sequences of catabolic genes encoding upper pathway of naphthalene biodegradation were reported for several Pseudomonas strains. They are: nah genes (for naphthalene degradation) from P. putida G7, NCIB 9816-4, BS202 and P. stutzeri AN10, dox genes (for dibenzothiophene oxidation) from Pseudomonas sp. C18, pah genes (for phenanthrene degradation) from P. putida OUS82 and P. aeruginosa PaK1. The gene organization and sequence similarity (about 90%) among the upper catabolic pathway genes of these strains were similar to those of the nah genes from the NAH7 plasmid. These genes are usually called ‘‘classical nah-like genes.’’

5.2.1

Naphthalene Catabolic Genes in P. stutzeri AN10 and Pseudomonas sp. ND6

Catabolic genes of the upper operon in mentioned above strains are arranged in the same order, with the exception of a nahQ-like gene deletion in P. stutzeri AN10. Two genes, encoding two independent salicylate 1-hydroxylases, nahG and nahW, have been found in P. stutzeri AN10. The gene nahG is a component of the same transcriptional unit that contains the genes for meta-cleavage catechol oxidation, forming the naphthalene degradation lower pathway. The gene, encoding NahW is situated outside but in close proximity (less than 3 kb) this transcriptional unit (Bosch et al., 1999). Both genes are induced by salicylate, and their products, NahG and NahW exhibit broad substrate specificity. However, 3-chlorosalicylate is better converted by NahW, whereas NahG is more efficient metabolizing methylsalicylates. Regulatory gene encoding the NahR-type protein is located between nahG and nahW genes. The plasmid pND6-1, 102 kb in size, isolated from naphthalene-degrading Pseudomonas sp. strain ND6, contains two duplicate naphthalene catabolic genes. One of them was identified as nahU (isofunctional gene of the classical salicylate hydroxylase gene), encoding a new salicylate hydroxylase NahU which possesses a higher binding ability to salicylate and cofactors and catalytic efficiency in comparison with NahG (Zhao et al., 2005). In pND6-1 and pDTG1 gene nahR is located upstream of nah2 operon, and both operons are transcribed in opposite direction towards each other. In NAH7, the regulatory gene nahR is located between nah1 and nah2 operons and the direction of transcription of the two operons is identical. The intervening region between the upper and lower nah operons of NAH7 does not show any homology with the corresponding regions of pDTG1 and pND6-1 plasmids. These data suggest that the nah operons on NAH7 have been acquired differently from those of pDTG1 and pND6-1 (Sota et al., 2006).

5.2.2

PAH Catabolic Genes Organization in P. putida BS202

In naphthalene and salicylate-degrading P. putida BS202 plasmid NPL-1 controls only primary stages of naphthalene degradation to salicylate. Salicylate oxidation to catechol with further ortho-cleavage is determined by chromosomal genes. However, the plasmid NPL-1 contains silent genes for meta-pathway and nonfunctioning nahG gene highly homologous to nahG from archetypal plasmid NAH7 (> Fig. 1). Remarkably, there is not any homology between

Diversity of Naphthalene Biodegradation Systems in Soil Bacteria

24

. Figure 1 Polycyclic aromatic hydrocarbons biodegradation by Pseudomonas putida BS202.

the chromosomal gene nahG1 and plasmid gene nahG. Comparison of the structure of NPL-1 (inducible synthesis of naphthalene dioxygenase) and NPL-41 (constitutive synthesis) revealed that an inverted DNA of 4.2 kb located upstream to the nah1-operon or overlapped may participate in the regulation of nah genes expression. The plasmid NPL-1 contains at least two transposons of the Tn3 family. These transposons are involved in rearrangements (deletions and inversions), and influence the expression of the catabolic and regulatory genes. The formation of a strong NahR-independent constitutive promoter by the inversion of a DNA fragment may be responsible for changing inducible synthesis of naphthalene dioxygenase to constitutive one. Change of inducible synthesis of naphthalene dioxygenase to constitutive one in strains resulted in acquisition the ability to degrade three-rings PAH, such as phenanthrene. Switching-on of the meta-pathway genes expands the range of potentially utilizable aromatic compounds and makes it possible strains to grow on methylated PAH which constitutes the major part of PAH in nature.

5.2.3

Organization of Naphthalene Catabolic Genes in P. putida AK5

Structural organization of new PAH biodegradative operon encoding oxidation of salicylate through gentisate in the strain P. putida АК5 has been studied (Izmalkova et al., manuscript in preparation). Genes for the catabolism of naphthalene and salicylate in P.putida strain AK5 are located on non conjugative IncP-7 plasmid pAK5, 135 kb in size. Utilization of naphthalene

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. Figure 2 Gene organization of the region of pAK5 plasmid in the strain P.putida AK5 encoding degradation of naphthalene via gentisic acid.

trough gentisate is performed by two catabolic operons. Both of them have their own regulatory gene (> Fig. 2). Transformation of naphthalene into salicylate is controlled by ‘‘classical’’ nah1-operon. Further degradation of salicylate is encoded by new sgp-operon including six ORFs – sgpAIKGHB. Four ORFs encode subunits of salicylate 5-hydroxylase: reductase component (sgpA), large subunit of oxygenase component (sgpG), small subunit of oxygenase component (sgpH) and ferredoxin (sgpB). Two ORFs for gentisate 1,2-dioxygenase (sgpI) and fumarylacetoacetate hydrolase (sgpK) were located between sgpA and sgpG in gene cluster encoding salicylate 5-hydroxylase. The gene for maleylpyruvate isomerase (mpi) has been localized downstream of sgp-operon. Organization of sgp-operon except for rearrangement of sgpI and sgpK genes, is similar to organization of catabolic genes in salicylate degradation operon in the plant pathogenic strains such as Ralstonia solanacearum GMI1000. However, cluster analysis did not reveal close relationship between them. Phylogenetic analysis of deduced amino acid sequences and order of the genes in sgp-operon indicate on mosaic structure of this operon.

5.3

Naphthalene Catabolic Genetic Systems Differ from Archetype Plasmid NAH7

Existence of highly homologous genes for the degradation of polycyclic aromatic hydrocarbons does not reflect metabolic diversity and potential of microorganisms. Many PAHdegrading microorganisms have no genetic relatedness to the ‘‘classical’’ nah genes described for Pseudomonas strains. Such bacteria as Comamonas sp., Burkholderia sp., Ralstonia sp. and some others contain genes for naphthalene and phenanthrene degradation that are different from those in bacteria belonging to genus Pseudomonas.

5.3.1

PAH Catabolic Genes of Comamonas testosteroni Strains

Three Comamonas testosteroni strains GZ38A, GZ39 and GZ42 capable of degrading naphthalene and/or phenanthrene were studied. The strain GZ39 possesses a new novel set of genes designated as the phd genes for naphthalene/phenanthrene degradation different from those found in P. putida strains. The phd genes for phenanthrene degradation in C. testosteroni GZ38A are similar but not identical to those in GZ39. The three Comamonas testosteroni strains represent at least two new classes of genes involved in PAH degradation. The phd genes order in C. testosteroni strain GZ39 is quite different from that determined for the nah genes found in Pseudomonas species. Several genes have not been identified in C. testosteroni strain GZ39, for example analog of nahC gene for 1,2-dihydroxynaphthalene dioxygenase.

Diversity of Naphthalene Biodegradation Systems in Soil Bacteria

24

The genes for naphthalene and phenanthrene degradation in C. testosteroni strain GZ42 were not the same but were similar to the classical nah genes of P. putida NCIB 9816. At least two new genes were found in the C. testosteroni GZ42 catabolic operon. Genes designated nahAc2 and nahAd2 are located between the genes nahAa and nahAb (Zylstra et al., 1997).

5.3.2

Naphthalene Catabolic Genes of Ralstonia sp. Strain U2

The operon for naphthalene biodegradation was studied in details in Ralstonia sp. strain U2 (Zhou et al., 2002). Ralstonia sp. strain U2 converts naphthalene to central metabolites via gentisate. There is similarity between the nag genes of strain U2 and the classical nah genes, but only in the conversion of naphthalene to salicylate. However, there are major differences: between nagAa and nagAb two genes (nagGH) were inserted. The genes nagG and nagH were very similar to nahAc2 and nahAd2 of strain C. testosteroni GZ42. NagG and NagH are structural subunits of salicylate 5-hydroxylase linked to electron transport proteins consisting of NagAb and NagAa. A further difference between nag and nah (NAH7 plasmid) clusters is in the location of the nagR and nahY. The genes for the complete pathway from naphthalene to pyruvate and fumarate appear to be on a single large operon spanning an 18-kb region, in which the genes for conversion of gentisate are directly downstream of the genes for conversion of naphthalene to salicylate.

5.3.3

Phenanthrene Catabolic Genes of Burkholderia sp. Strain RP007

Naphthalene and phenanthrene are degraded by Burkholderia sp. RP007 through common routes via salicylate and 1-hydroxy-2-naphthoic acid, respectively (Laurie and Lloyd-Jones, 1999). The phn genes of Burkholderia sp. RP007 constitute a plasmid-borne locus which encodes an upper pathway for PAH catabolism that has different gene order and that contains isofunctional genes of low homology in comparison to the classical nah-like genes. The locus of initial dioxygenase contains genes phnAc and phnAd but lacks both the ferredoxin and reductase components. The phnB gene encoding cis-diol dehydrogenase is more closely related to the corresponding genes from biphenyl catabolic pathways than to those described for classical nahB genes. Another gene, nahC encoding PAH extradiol dioxygenase shows a phylogeny not seen before among extradiol dioxygenases from any PAH or biphenyl catabolic pathway. Two catechol 2,3-dioxygenase genes have been characterized for strain RP007. Two putative regulatory genes, phnR and phnS were found upstream of phn catabolic genes. The phnS is a LysR-type transcriptional activator and is cotranscribed with catabolic genes as a part of an operon. The gene phnR encodes a s54-dependent regulator. It was suggested that that phn locus was compiled by the recruitment of individual genes from a variety of catabolic pathways.

5.3.4

Degradation of PAH by Sphingomonas sp

The members of the genus Sphingomonas and related species are capable of growth on both monocyclic and polycyclic hydrocarbons. For example, Novosphingobium aromaticivorans strain F199 can grow on toluene, isomers of xylene, p-cresol, biphenyl, naphthalene,

1161

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Diversity of Naphthalene Biodegradation Systems in Soil Bacteria

dibenothiophene, fluorine, salicylate, and benzoate (Habe and Omori, 2003). At least 13 gene clusters were predicted to encode enzymes involved in degradation of aromatic compounds in this strain and were complexity arranged in pNL1, a 184-kb catabolic plasmid. Seven sets of oxygenase components seemed to interact with only set of ferredoxin and reductase components in plasmid pNL1. The results obtained suggested that unusual arrangement of various genes from different catabolic pathways may be typical of Sphingomonas species.

5.3.5

Naphthalene Catabolic Genes of Rhodococcus sp

Clusters of genes which include determinants for the catalytic subunits of naphthalene dioxygenase (narAa and narAb) were analyzed in naphthalene-degrading Rhodococcus strains (Larkin et al., 1999). In the region analyzed homologous gene clusters are separated from each other by non homologous DNA, and there are various degrees of homology between related genes, and that nar genes are located on plasmids in strains NCIMB12038 and P400 and on a chromosome in P200. The nar region is not organized into a single operon but there are several transcription units which differ in the strains investigated. The narA and narB genes were found to be transcribed as a single unit in all strains analyzed, and their transcription was induced by naphthalene. The putative aldolase gene (narC) was found on the same transcript only in strains P200 and P400. Putative regulatory genes (narR1 and narR2) were transcribed as a single mRNA in naphthalene-induced cells. A number of the genes known to be essential for naphthalene catabolism in Gram-negative bacteria were not found in the region analyzed.

6

Variability of PAH Catabolic Systems in Naturally Occurring Bacteria

As mentioned above the relative position of the catabolic operons, structural and regulatory genes vary. Naphthalene operons can be located in trans-position as it has been shown for P. putida strains bearing NPL-1, pBS1141 and pBS1191. In the first two cases nah1 operon is located on the plasmids, and nah2 operon – on the host bacterial chromosome, in the third case gene encoding key enzyme of nah2 operon (salG) is located on another plasmid – pBS1192. New groups of key naphthalene biodegradative genes, nahAc and nahG, were found (Izmalkova et al., 2005). Thus, nah-like genetic systems, despite of conservative organization and high level of homology demonstrate significant variability. Catabolic pathways are continuously subjected to selective pressure and various combinations of catabolic genes in different Pseudomonas sp. strains have arisen over time. Genetic plasticity of catabolic operons plays a significant role in adaptation of microorganisms to a wide variety of environmental conditions.

7

Research Needs

It has been shown recently that many NAH-like plasmids belong to unknown incompatibility groups. Study of basic replicons of new isolated plasmids is necessary for classification of degradative plasmids. It is not known why IncP-9 degradative plasmids are widely distributed in different soils while IncP-7 plasmids could be found mainly in highly polluted soils. In order

Diversity of Naphthalene Biodegradation Systems in Soil Bacteria

24

to assess the prevalence of different PAH-catabolic genotypes in nature and to follow up the evolution of modern catabolic pathways there is a need for more complete nucleotide sequences for plasmids and degradative operons found in different PAH-degrading microorganisms.

References Boronin AM (1992) Diversity of Pseudomonas plasmids: to what extent? FEMS Microbiol Lett 100: 461–468. Bosch R, Moore ERB, Garcia-Valdes E, Pieper DH (1999) NahW, a novel, inducible salicylate hydroxylase involved in mineralization of naphthalene by Pseudomonas stutzeri AN10. J Bacteriol 181: 2315–2322. Habe H, Omori T (2003) Genetics of polycyclic aromatic hydrocarbon metabolism in diverse aerobic bacteria. Biosci Biotechnol Biochem 67: 225–243. Izmalkova TYu, Sazonova OI, Sokolov SL, Kosheleva IA, Boronin AM (2005) Diversity of genetic systems responsible for naphthalene biodegradation in Pseudomonas fluorescens strains. Microbiology (Russian) 74: 60–68. Larkin MJ, Allen CC, Kulakov LA, Lipscomb DA (1999) Purification and characterization of a novel naphthalene dioxygenase from Rhodococcus sp. strain NCIMB12038. J Bacteriol 181: 6200–6204. Laurie AD, Lloyd-Jones G (1999) The phn genes of Burkholderia sp. strain RP007 constitute a divergent gene cluster for polycyclic aromatic hydrocarbons catabolism. J Bacteriol 181: 531–540. Park W, Jeon CO, Madsen EL (2002) Interaction of NahR, a LysR-type transcriptional regulator, with the alpha subunit of RNA polymerase in the naphthalene degrading bacterium, Pseudomonas putida NCIB 9816–4. FEMS Microbiol Lett 213: 159–165. Sevastsyanovich YR, Krasowiak R, Bingle LEH, Haines AS, Sokolov SL, Kosheleva IA, Leuchuk AA,

Titok MA, Smalla K, Christopher M, Thomas CM (2008) Diversity of IncP-9 plasmids of Pseudomonas. Microbiology 154: 2929–2941. Shell MA (1990) Regulation of the naphthalene degradation genes of plasmid NAH7: example of a generalized positive control system in Pseudomonas and related bacteria. In Pseudomonas: Biotransformation, Pathogenesis, and Evolving Biotechnology. Washington, DC: American Society for Microbiology, pp. 165–176. Sota M, Yano H, Ono A, Miyazaki R, Ishii H, Genka H, Top E, Tsuda M (2006) Genomic and functional analysis of the IncP-9 naphthalene-catabolic plasmid NAH7 and its transposon Tn4655 suggests catabolic gene spread by a tyrosine recombinase. J Bacteriol 188: 4057–4067. Zhao H, Chen D, Li Y, Cai B (2005) Overexpression, purification and characterization of a new salicylate hydroxylase from naphthalene-degrading Pseudomonas sp. strain ND6. Microbiol Res 160: 307–313. Zhou N-Yi, Al-Dulayymi J, Baird MS, Williams PA (2002) Salicylate 5-hydroxylase from Ralstonia sp. strain U2: a monooxygenase with close relationships to and shared electron transport proteins with naphthalene dioxygenase. J Bacteriol 184: 1547–1555. Zylstra GJ, Kim E, Gloyal AK (1997) Comparative molecular analysis of genes for polycyclic aromatic hydrocarbon degradation. Genet Eng 19: 257–269.

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25 Genomic View of Mycobacterial High Molecular Weight Polycyclic Aromatic Hydrocarbon Degradation O. Kweon . S.-J. Kim . C. E. Cerniglia* Division of Microbiology, National Center for Toxicological Research, Food and Drug Administration, Jefferson, AR, USA *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1166

2 Bacterial Metabolism of HMW PAHs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1167 2.1 Oxygenases as Key Enzymes in the Degradation of HMW PAHs . . . . . . . . . . . . . . . . . . 1167 2.2 Multiple Pathways for the Degradation of HMW PAHs . . . . . . . . . . . . . . . . . . . . . . . . . . . 1168 3 Molecular Background of HMW PAH Degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1170 3.1 Genomic Insights into the Degradation of HMW PAHs . . . . . . . . . . . . . . . . . . . . . . . . . . . 1170 3.2 Regulation of Genes Involved in the Degradation of HMW PAHs . . . . . . . . . . . . . . . . 1173 4

Research Needs and Future Directions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1175

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_81, # Springer-Verlag Berlin Heidelberg, 2010

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Genomic View of Mycobacterial HMW PAH Degradation

Abstract: Bacteria play an important role in the degradation of high-molecular-weight (HMW) polycyclic aromatic hydrocarbons (PAHs), which are recalcitrant in the environment and toxic to living organisms. A diverse bacterial community has been identified and characterized from PAH-contaminated soils and sediments. Strains of mycobacteria have most frequently been found to degrade HMW PAHs (> Chapter 19, Vol. 3, Part 1). The nidAB- and nidA3B3-like oxygenase genes of these strains have been reported to be involved in the initial ring-hydroxylation of HMW PAHs. There are significant differences between the genetic systems in mycobacteria and other bacterial strains that are widely known to degrade monocyclic aromatic compounds or low-molecular-weight (LMW) PAHs. Genes involved in PAH degradation are organized in an atypical mosaic structure made of a complex arrangement of gene clusters. Recent functional genomic analyses of mycobacteria have also identified a high number of catabolic genes with a wide range of substrate specificities and multiple pathways for the degradation of HMW PAHs. Multiple gene copies encoding Rieske nonheme ringhydroxylating oxygenases (21 copies) and cytochrome P450 monooxygenases (50 copies) were identified from Mycobacterium vanbaalenii PYR-1. Systems biology techniques are being applied to provide global insights into the genes, enzymes, and the regulatory mechanisms for HMW PAH degradation.

1

Introduction

The fate in nature of high-molecular-weight (HMW) PAHs with four or more fused aromatic rings is of great concern because of their genotoxic, mutagenic, and carcinogenic effects and their accumulation in the food chain, which poses significant risks to the environment and living organisms (Cerniglia, 1992). There are a number of environmental factors affecting microbial metabolism of HMW PAHs, which lead to their persistence in soil. Inherent recalcitrance due to their stable physico-chemical structures also hampers the microbial degradation of HMW PAHs (Cerniglia, 1992; Kanaly and Harayama, 2000). Adaptation strategies and PAH bioavailability-promoting mechanisms exist to overcome these obstacles. These include low energy or oxygen requirements for cell maintenance, a high specific affinity for PAH substrates (Fritzsche, 1994; Wick et al., 2002), cell surface hydrophobicity or modification of bacterial cell wall composition (Wick et al., 2003b), multiple substrate utilization (Wick et al., 2003a), capacity to form biofilms (Bastiaens et al., 2000; Child et al., 2007; Wick et al., 2002), biosurfactant production (Das and Mukherjee, 2007; Johnsen et al., 2005), and even surface motility (Fredslund et al., 2008). However, from a fundamental standpoint, HMW PAH degradation strategies involve the function of versatile catabolic genes and enzymes as well as their efficient regulation (Kim et al., 2008). Thorough reviews on the environmental factors and biological strategies for the microbial degradation of PAHs have been written (Cerniglia and Sutherland, 2006; Das and Mukherjee, 2007; Juhasz and Naidu, 2000; Johnsen et al., 2005; Kanaly and Harayama, 2000; Van Hamme et al., 2003). Genetic and biochemical mechanisms for the degradation of aromatic compounds consisting of three or fewer benzene rings have been reviewed (Andreoni and Gianfreda, 2007; Cerniglia, 1992; Habe and Omori, 2003; Diaz, 2004; Johnsen et al., 2005; Williams and Sayers, 1994). However, studies on the enzymes and catabolic genes involved in HMW PAH degradation are not numerous (Kanaly and Harayama, 2000). Relatively few bacterial strains have been isolated as HMW PAH-degraders, in comparison to the large number of bacteria capable of degrading LMW aromatic compounds, perhaps due to lower bioavailability of HMW PAHs

Genomic View of Mycobacterial HMW PAH Degradation

25

(Kanaly and Harayama, 2000). Although several Gram-negative strains of Pseudomonas and Sphingomonas have been reported to degrade HMW PAHs (Das and Mukherjee, 2007; Gibson et al., 1975; Kanaly and Harayama, 2000; Kazunga and Aitken, 2000; Pinyakong et al., 2003), most organisms degrading HMW PAHs are nocardioform Gram-positive actinomycetes, especially members of the genera Mycobacterium (Boldrin et al., 1993; Grosser et al., 1991; Heitkamp et al., 1988; Kelley and Cerniglia, 1995; Lo´pez et al., 2005; Schneider et al., 1996; Vila et al., 2001) (> Chapter 19, Vol. 3, Part 1) and Rhodococcus (Walter et al., 1991). Direct functional correlation of genes with the degradation of HMW PAHs has been demonstrated in Mycobacterium spp. (Khan et al., 2001; Kim et al., 2006; Krivobok et al., 2003; Pagnout et al., 2007). Molecular ecological studies have shown that genetic systems and metabolic processes for the degradation of HMW PAHs in mycobacterial strains are not closely related to their counterparts from other microorganisms (Habe and Omori, 2003; Kim et al., 2008). For example, primer- or probe-based approaches based on the sequence of nahAc- and phnAc-like genes from Pseudomonas, Sphingomonas, and Burkholderia spp. do not detect those genes in mycobacteria (Brezna et al., 2003; Churchill et al., 1999; Hall et al., 2005; Hamann et al., 1999; Zhou et al., 2006). These observations suggest either different origins for the aromatic catabolic genes from mycobacteria or significant genetic divergence from those from Gramnegative bacteria. In recent years, the knowledge concerning the physiology, biochemistry, and genetics with respect to HMW PAH metabolism has increased using ‘‘omics’’ methodologies, including advanced functional genomic techniques. These studies have shown that the members of the genus Mycobacterium appear to be specialized for the degradation of HMW PAHs (Kim et al., 2008; Leys et al., 2005). This review focuses on the bacterial degradation of HMW PAHs, with special emphasis on the molecular analysis of PAH-degrading mycobacteria in the environment.

2

Bacterial Metabolism of HMW PAHs

2.1

Oxygenases as Key Enzymes in the Degradation of HMW PAHs

The metabolism of HMW PAHs proceeds by mono- and dioxygenation reactions in bacterial systems. In this enzymatic step, either one or two atoms of dioxygen are incorporated into substrates, forming dihydrodiol compounds with trans and cis configurations, respectively (Kelley et al., 1990; Heitkamp et al., 1988). Two groups of enzymes, multicomponent Riesketype nonheme mono-iron ring-hydroxylating oxygenases (RHO) and heme cytochrome P450 monooxygenase (CYP), are critical to initiate the aerobic metabolism of HMW PAHs (Cerniglia, 1992). Oxygenases are key enzymes because they limit the range of compounds that can be degraded by microbial catabolic systems. The bacterial oxidation of HMW PAHs is mostly initiated by dioxygenation. Analysis of metabolites formed by RHOs revealed that these enzymes catalyze the oxidation of a structurally diverse range of PAHs. Recently, several genes encoding the oxygenase components of RHO enzymes from pyrene- and fluoranthene-degrading mycobacterial strains have been analyzed in detail with the illustration of their substrate specificities (Kim et al., 2006; Khan et al., 2001; Krivobok et al., 2003). Khan et al. (2001) reported that the products of pyreneinduced oxygenase nidAB genes from M. vanbaalenii PYR-1 catalyze the conversion of pyrene. The same research group showed that RHO nidA3B3 genes from the same bacterium are

1167

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Genomic View of Mycobacterial HMW PAH Degradation

upregulated by fluoranthene but not by pyrene. Both RHO systems are able to transform a wide range of PAHs (Kim et al., 2006; Khan et al., 2001; Krivobok et al., 2003). Similar RHO systems, Pdo1 and Pdo2 from Mycobacterium sp. strain 6PY1 and PhdAB from Mycobacterium sp. strain SNP11, have been identified (Krivobok et al., 2003; Pagnout et al., 2007). These genes show broad substrate affinities for benz[a]anthracene, fluoranthene, pyrene, phenanthrene, anthracene, fluorene, and dibenzofuran. The nid-, pdo-, and phd-like genes form a phylogenetically independent cluster distinct from those classical RHOs. They are found in Mycobacterium, Nocardioides, Rhodococcus, Terrabacter, and Arthrobacter spp. (Kweon et al., 2008). Another interesting feature of the bacterial degradation of HMW PAHs is the involvement of CYP enzymes, which have often been observed in fungal and mammalian systems (Cerniglia, 1992; Moody et al., 2004). CYP enzymes, along with epoxide hydrolase, are involved in the oxygenation of a wide range of organic molecules (Bernhardt, 2006; Cerniglia and Sutherland, 2006; Urlacher and Schmid, 2002). Although CYP enzyme activity appears to be lower than RHO activity (Cerniglia, 2003), functional studies have indicated that CYPs are responsible for the oxidation of PAHs, including naphthalene, phenanthrene, dibenzothiophene, fluoranthene, pyrene, 7-methylbenz[a]anthracene, and benzo[a]pyrene (Brezna et al., 2006; England et al., 1998; Harford-Cross et al., 2000; Taylor et al., 1999).

2.2

Multiple Pathways for the Degradation of HMW PAHs

One of the features in the bacterial degradation of HMW PAHs is the presence of multiple pathways for the degradation of the same PAH, although the substrate is usually converted into a limited number of central metabolic intermediates. This may result from multiple gene copies encoding various initial oxygenase enzymes. M. vanbaalenii initiates the degradation of pyrene and fluoranthene with at least two and four different degradation routes, respectively, most of which are linked to the TCA cycle via protocatechuate (Kim et al., 2005, 2007; Kweon et al., 2007). Pyrene has been used as a model compound to study biodegradation of HMW PAHs since it is structurally similar to several carcinogenic PAHs (Kanaly and Harayama, 2000). Studies have been conducted with various pyrene degraders, generally mycobacterial strains, to establish pathways of pyrene degradation based on identified metabolites, 18O2 incorporation experiments, and genetic/enzymatic mechanism information (Dean-Ross and Cerniglia, 1996; Heitkamp et al., 1988; Kim et al., 2005; Khan et al., 2001; Krivobok et al., 2003; Schneider et al., 1996; Vila et al., 2001; Walter et al., 1991). These efforts have provided the first detailed description of the bacterial pathway for the complete mineralization of pyrene into CO2 (Kim et al., 2007). Pyrene-degrading mycobacterial strains initially oxidize this PAH by two routes (> Fig. 1). The first, minor route, an initial dioxygenation at the C-1 and C-2 positions, forms an O-methylated derivative of pyrene-1,2-diol as a detoxification step (Kim et al., 2004). The second route is the predominant pathway, in which dioxygenation is initiated at the C-4 and C-5 positions of the K-region to give pyrene cis-4,5-dihydrodiol. Rearomatization of the dihydrodiol and subsequent ring cleavage dioxygenation leads to the formation of 4,5-dicarboxyphenanthrene, which is further decarboxylated to 4-phenanthroate. The subsequent intermediate, cis-3,4-dihydroxyphenanthrene-4-carboxylate, is produced by a second dioxygenation reaction. Rearomatization then forms 3,4-dihydroxyphenanthrene, which is further metabolized to 1-hydroxy-2-naphthoate. The next enzymatic reaction, including intradiol ring-cleavage dioxygenation, results in the production of o-phthalate.

Genomic View of Mycobacterial HMW PAH Degradation

. Figure 1 (Continued)

25

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25

Genomic View of Mycobacterial HMW PAH Degradation

Phthalate is then degraded via the b-ketoadipate pathway through protocatechuate and finally mineralized to CO2 via the TCA cycle (Heitkamp et al., 1988; Krivobok et al., 2003; Kim et al., 2005; Kim et al., 2007; Pagnout et al., 2007). Detoxification pathways are additional routes in HMW PAH degradation (Kim et al., 2003, 2007, 2008). This pathway does not involve carbon assimilation but is essential for bacterial cells to deal with potentially toxic compounds. For example, dihydrodiol dehydrogenases catalyze the conversion of PAH dihydrodiols to highly unstable dihydroxy-PAHs, which are then nonenzymatically oxidized to o-quinones (Penning et al., 1999). These dihydroxy-PAHs and o-quinones are biologically active and are known to be deleterious to cells (Penning et al., 1999). Catechol O-methyltransferases and o-quinone reductases found in PAH-degrading mycobacterial strains are involved in neutralization of the potential toxic effects of these PAH metabolites (Kim et al., 2003, 2004).

3

Molecular Background of HMW PAH Degradation

3.1

Genomic Insights into the Degradation of HMW PAHs

Recently, the genomes of five pyrene-degrading mycobacterial isolates have been completely sequenced and interesting features involved in the degradation of PAHs have emerged. Kim et al. (2007) analyzed one of these environmental isolates, M. vanbaalenii PYR-1, with respect to the degradation of pyrene using a combination of metabolic, genomic, and whole-cell proteomic approaches, in which 27 functional enzymes necessary for the complete degradation of pyrene were identified (> Fig. 1 and > Table 1). They precisely correlated the pyrene degradation pathway with the enzyme abundance for each metabolic step. The metabolism of fluoranthene in M. vanbaalenii was also investigated using several polyomic approaches (Kweon et al., 2007). Fifty-four enzymes involved in the degradation of fluoranthene were reported. A comprehensive picture of the overall molecular basis for the bacterial metabolism of HMW PAHs now has been provided from the further analysis of complete genome sequence of M. vanbaalenii (Kim et al., 2007; Kim et al., 2008; Kweon et al., 2007). The 6.5 Mb genome of M. vanbaalenii contains 5,979 predicted protein coding sequences in a single circular chromosome with an average G + C content of 67%. Analysis of the genome has . Figure 1 Complete pyrene degradation pathway in M. vanbaalenii PYR-1 based on metabolite, genome, and proteome analyses (adapted from Kim et al., 2007). The letter ‘‘S’’ denotes enzymatic reaction steps (> Table 1). Pyrene metabolic intermediates are as follows: P1, pyrene cis-4,5-dihydrodiol; P2, 4,5-dihydroxypyrene; P3, phenanthrene-4,5-dicarboxylate; P4, phenanthrene-4-carboxylate; P5, cis-3,4-dihydroxyphenanthrene-4-carboxylate; P6 3,4-dihydroxyphenanthrene; P7, 2-hydroxy-2H-benzo[h]chromene-2-carboxylate; P8, 1-hydroxy-2-naphthaldehyde; P9, 1-hydroxy-2-naphthoate; P10, trans-2’carboxybenzalpyruvate; P11, 2-carboxybenzaldehyde; P12, phthalate; P13, phthalate 3,4-dihydrodiol; P14, 3,4-dihydroxyphthalate; P15, protocatechuate; P16, b-carboxy-cis, cis-muconate; P17, g-carboxymuconolactone; P18, b-ketoadipate enol-lactone; P19, b-ketoadipate; P20, b-ketoadipyl CoA; P21, pyrene cis-1,2-dihydrodiol; P22, 1,2dihydroxypyrene; P23, 1-methoxy-2-hydroxypyrene; P24, 1-hydroxy-2-methoxypyrene; P25, 1,2-dimethoxypyrene; P26, pyrene o-quinone.

25

Genomic View of Mycobacterial HMW PAH Degradation

. Table 1 Proteins identified in the whole-cell proteome of M. vanbaalenii PYR-1 grown in the presence of pyrene Metabolic enzymatic reactiona

Gene/ (Mvan ID)b

S1

nidA/ 0488

Pyrene ring-hydroxylating oxygenase, a subunit

PdoA1

98

Mycobacterium sp. 6PY1

S1

nidB/ 0483

Pyrene ring-hydroxylating oxygenase, b subunit

PdoB1

98

Mycobacterium sp. 6PY1

S2/6/23

0544

Dihydrodiol dehydrogenase

PhdE

76

Nocardioides sp. KP7

S3/7

phdF/ 0470

Ring-cleavage dioxygenase

PhdF

83

Nocardioides sp. KP7

S4/15

0543

Decarboxylase

PhtC

74

Arthrobacter keyseri 12B

S5

0546

Phenanthrene ringhydroxylating oxygenase, a subunit

PdoA2

99

Mycobacterium sp. 6PY1

S5

0547

Phenanthrene ringhydroxylating oxygenase, b subunit

PdoB2

99

Mycobacterium sp. 6PY1

S8

phdG/ 0472

Hydratase-aldolase

PhdG

84

Nocardioides sp. KP7

S9

nidD/ 0486

Aldehyde dehydrogenase

PhdH

85

Nocardioides sp. KP7

S10

phdI/ 0468

1-Hydroxy-2-naphthoate dioxygenase

PhdI

46

Nocardioides sp. KP7

S11

phdJ/ 0469

trans-2’Carboxybenzalpyruvate hydratase-aldolase

PhdI

55

Nocardioides sp. KP7

S12

0522

2-Carboxylbenzaldehyde dehydrogenase

PhdK

60

Nocardioides sp. KP7

S13

phtAa/ Phthalate 3,4-dioxygenase, a subunit 0463

PhtA1

74

Terrabacter sp. DBF63

S13

phtAb/ Phthalate 3,4-dioxygenase, 0464 b subunit

PhtA2

68

Terrabacter sp. DBF63

S1/5/13/22

phtAc

Oxygenase ferredoxin component

PhtAc

69

Arthrobacter keyseri 12B

S1/5/13/22

phtAd/ Oxygenase reductase 0467 component

PhtAd

59

Arthrobacter keyseri 12B

S14

0466

PhtB

64

Terrabacter sp. DBF63

Matching protein Enzyme

Phthalate 3,4-dihydrodiol dehydrogenase

Proteinc Similarityd Source organism

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Genomic View of Mycobacterial HMW PAH Degradation

. Table 1 (Continued) Matching protein

Metabolic enzymatic reactiona

Gene/ (Mvan ID)b

S16

0561

Protocatechuate 3,4dioxygenase, a subunit

PcaG

45

Streptomyces sp. 2065

S16

0560

Protocatechuate 3,4dioxygenase, b subunit

PcaH

58

Streptomyces sp. 2065

S17

0562

b-Carboxy-cis,cis-muconate cycloisomerase

PcaB

44

Terrabacter sp. DBF63

S18/19

0563

g-Carboxymuconolactone decarboxylase/b-ketoadipate enol-lactone hydrolase

PcaL

40

Rhodococcus opacus 1CP

S20

0564

b-Ketoadipate succinyl CoA transferase, a subunit

PcaI

64

P. putida PRS2000

S20

0565

b-Ketoadipate succinyl CoA transferase, b subunit

PcaJ

60

P. putida PRS2000

S21

4589

b-Ketoadipyl CoA thiolase

PcaF

36

Terrabacter sp. DBF63

S22

nidA3/ 0525

Ring-hydroxylating oxygenase, PdoA a subunit

97

Terrabacter sp. HH4

S22

nidB3/ 0526

Ring-hydroxylating oxygenase, PdoB b subunit

98

Terrabacter sp. HH4

S24/25

COMT/ Catechol O-methyltransferase 3280

MT1743

72

M. tuberculosis CDC1551

S26

PQR/ 2039

Quinone reductase

MonD

45

S. cinnamonensis ATCC15413

0600

Cytochrome P450 monooxygenase

Ema7

66

Streptomyces sp. HIS-0435

3012

Cytochrome P450 monooxygenase

MonD

45

S. cinnamonensis ATCC15413

3029

Cytochrome P450 monooxygenase

MonD

45

S. cinnamonensis ATCC15413

3108

Cytochrome P450 monooxygenase

SpiL

33

Sorangium cellulsum Soce90

CYP51/ Cytochrome P450 5161 monooxygenase

CYP51

66

Solanum chacoense

0521

Ephx1

36

Oryctolagus cuniculus

Enzyme

Epoxide hydrolase I

Proteinc Similarityd Source organism

Indicates pyrene metabolic steps in > Fig. 1 ORF (Mvan ID) indicates the locus tag number assigned to each ORF in the M. vanbaalenii PYR-1 complete sequences. If no number listed, no ORF was identified in the genome for the corresponding protein b,c Gene/protein in boldface type was functionally characterized d Percent identity was based on alignments with BlastP hits from the nonredundant NCBI protein database a

b

Genomic View of Mycobacterial HMW PAH Degradation

25

revealed 194 chromosomally encoded genes that are likely associated with the degradation of aromatic compounds. The genes and their functional annotations are described in detail by Kim et al. (2008). The most distinctive feature of the genome is a 150 kb major catabolic island at positions 494–643 kb (region A) with an additional 31 kb catabolic island at positions 4,711–4,741 kb (region B). These are predicted to encode most of the enzymes for the degradation of PAHs. The 150 kb catabolic region A appears to be specialized in the degradation of HMW PAHs, since it possesses all the catabolic genes required for complete PAH degradation, including those of the pyrene and the b-ketoadipate pathways (Kim et al., 2008). Significant differences in the gene structure and organization from other well-characterized aromatic hydrocarbon degraders, including Pseudomonas, Burkholderia, Sphingomonas, and Rhodococcus were revealed. The catabolic genes in pseudomonads and closely related genera are usually well organized in a cluster (Assinder and Williams 1990; van der Meer et al., 1992). However, the organization of catabolic genes in mycobacteria has an atypical mosaic pattern made of several complex gene clusters. Genes involved in a degradative pathway are not arranged on the same operon but are dispersed throughout several gene clusters. The 27 genes involved in the complete degradation of pyrene are scattered over at least 7 operons, which are positioned across 4 gene clusters (Kim et al., 2008). For example, genes Mvan_0487/0488, Mvan_0544, and Mvan_0470, encoding the first three steps of pyrene degradation (RHO, cis-dihydrodiol dehydrogenase, and ring-cleavage dioxygenase), are located in three different clusters over 70 kb of region A (Kim et al., 2008). Genes Mvan_0467/phtAc, coding for the electron transfer components of the initial pyrene dioxygenase, are not even found together with the oxygenase components in the same gene cluster. Many identified genes are enriched with multiple paralogs, showing a remarkable range of diversity (Kim et al., 2008). A significantly high number of gene copies encoding RHOs (21 copies) and CYPs (50 copies) have been identified in the genome of M. vanbaalenii (> Fig. 2). The redundancy of genes, which also includes genes other than oxygenases, is thought to contribute to the versatile PAH degradation capacity of M. vanbaalenii (Kim et al., 2007; Kweon et al., 2007; Kim et al., 2008). High similarity in both sequence and gene orientation has been reported among PAHdegrading mycobacterial catabolic genomic regions, including nidAB and other genes involved in pyrene degradation (Brezna et al., 2003; Krivobok et al., 2003; Habe et al., 2004; Miller et al., 2004; Sho et al., 2004; Johnsen et al., 2006; Pagnout et al., 2007). This finding suggests that these genetic systems are evolutionarily related and play an important role in the transformation of HMW PAHs. A number of transposase-like genes are commonly identified in the vicinity of PAH catabolic gene clusters (Sho et al., 2004; Kim et al., 2008). These multiple mobile genetic elements are likely involved in the transfer of catabolic modules between bacteria or in the duplication or rearrangement of genes within the same bacterium (Habe and Omori 2003; Pinyakong et al., 2003; Sho et al., 2004).

3.2

Regulation of Genes Involved in the Degradation of HMW PAHs

Little is known about the regulation of genes and enzymes involved in the degradation of HMW PAHs. Genetic elements have not been directly correlated to regulatory mechanisms. Pyrene metabolism is inducible in M. vanbaalenii (Heitkamp and Cerniglia 1988; Cerniglia 2003). Functional genomic analysis has demonstrated that 18 enzymes in the pyrene degradation pathway are upregulated more than twofold (Khan et al., 2001; Kim et al., 2007). In another study, performed with M. vanbaalenii exposed to fluoranthene, some enzymes are also

1173

25

Mvan 0533

Mvan 0539 100 546 100 Mvan 0 84 0 KP7 10 phdA 25 100 100 n 05 1 Mva 6PY A1 88 Pdo 04 5 an 89 S6 Mv A oA 12 Pd dA F63 Ni DB 1

tA ph

pa

dA

tA

a

va

n

04

B

94

AB

100

a2 6 RH 3 A1 K Op ME 84 hA -9 2D 10 BO 0 Oph 1 A2 D B01 1 Pht3 00 putid 100 a cbaA BR60 100 pobA POB3 88 10

Dit

0 10

Mvan 1001

VanA HR199

100

100

NagAc U2

100 100

100

PhnA1

b CHY -1 AhdA1c P2 BPhA1d F199

9 F19

A1c

Bph

Mva

n4

421 3 910

90 84

an

Mv

an

41

41

Mv

100

10

62

49

19

an

n

43

n Mv

nahAc 9816-4 100 pahAc OUS8 2 atdA 100 YAA t d 100 nA1 ucc OR 22 F7 10 NC Ps 0 7N bA cN Cm o-7 an tAb F t 1 An A A D tA CA P1 10

P2 9 lX 19 Xy lX F BS Xy A 2C P1 bd 0 AD c 10 nA 2 Be 049 an 100 v M P2 A1e Ahd 199 1e F BphA 100 100 U2 NagG 100 HybB JB2 AhdA1d P2

0

10 0 100

va

98

100

M

100

10

0

100

100

tsaM T-2 100 0 10 5211 Mvan 100 5 2 2 5 n Mva 0 10 69 n 28 0 Mva 10 012 2 an 84 Mv 7 3 an 10 Mv CA -9 a 4 A r DN 08 ca XL 09 a n rA va a M C va

01

IP

99 F1 m b u C hA1 P2 Bp A1b 0 0 d P2 1 Ah 1a dA 199 h A aF A1 h 0 2 p B 10 AFK Ac 0 phn 10 1 A5 PhnA 81 f F199 BphA1 100 100 HY-1 PhnA1a C 99 PahA3 pak1

A1

12

92

ph

M

dxnA1 RW1 carAa CB 3 100 BphA 96 1 TA4 21 100 Mva n 44 15 10 10 b 0 p 0 hA1 RHA tod 1 10 C1 0 F1 Tc bA aP 90 bp 51 h bp A L B4 h bp A1 00 hA KF 1 KK 707 S1 02

Genomic View of Mycobacterial HMW PAH Degradation

M

1174

. Figure 2 Cladogram of 21 ring-hydroxylating oxygenases of M. vanbaalenii obtained from alignment with 59 related proteins from other microorganisms. The M. vanbaalenii genes are in red. The numbers on branches refer to the percentage confidence, estimated by a bootstrap analysis with 100 replications.

commonly induced (Kweon et al., 2007). Since many reactions and metabolic products in PAH metabolism are typical of the degradation pathway or are frequently connected within pathways, it is reasonable for M. vanbaalenii to share enzymes and reaction steps for the degradation of PAH substrates. For example, enzymes for protocatechuate degradation (e.g., protocatechuate 3,4-dioxygenase) are always detected in cells growing on pyrene or fluoranthene (Kim et al., 2007; Kweon et al., 2007). The expression of enzymes involved in certain steps of PAH degradation is significantly affected by growth in the presence of different PAH sources, as revealed by the changes in proteome abundance. For example, the oxygenase components, NidAB and NidA3B3, are highly upregulated only in the presence of pyrene and fluoranthene, respectively, indicating that they are subjected to different regulation mechanisms (Kim et al., 2007; Kweon et al., 2007). However, it is not clear if the mode of

Genomic View of Mycobacterial HMW PAH Degradation

25

regulation in M. vanbaalenii for the degradation of pyrene and fluoranthene is a sequential induction pattern. In sequential induction, enzymes are only synthesized when the respective metabolite is formed upon degradation of aromatic compounds, as in several pseudomonads (Assinder and Williams 1990; van der Meer et al., 1992; Ramos et al., 1997). Since genes responsible for the degradation pathways of mycobacteria are not organized in such operonic structures as are found in pseudomonads, the regulation of PAH-degrading genes may require several specific regulatory proteins to respond to different substrates or degradation intermediates (Krivobok et al., 2003). Sequence analysis of the M. vanbaalenii genome has identified multiple genes potentially encoding transcriptional regulators (Kim et al., 2007; Kweon et al., 2007; Kim et al., 2008).

4

Research Needs and Future Directions

Both HMW PAH ring-hydroxylating enzymes, NidAB and NidA3B3, from M. vanbaalenii show relaxed oxygenation activities (Khan et al., 2001; Kim et al., 2006). These results could suggest that both enzymes might be involved in the initial attack on pyrene or fluoranthene in M. vanbaalenii. However, the difference between responsiveness of these enzymes upon exposure to pyrene and fluoranthene indicates that in M. vanbaalenii the initial attack of each PAH substrate involves NidAB for pyrene and NidA3B3 for fluoranthene, although both enzymes have overlapping substrate specificities. This interpretation required high-throughput analytical genomic technologies to be performed. Genomic methodologies provide substantial benefits for understanding the mechanism of regulation that otherwise would be unpredictable. However, the outline of regulatory mechanisms of microbial physiology with respect to HMW PAH degradation is still not clear. Considering the structural complexity of the genes involved in the degradation of HMW PAHs, a number of factors are likely to influence the expression of catabolic genes to accommodate the metabolic and physiological needs of bacteria. The function of each of these factors needs to be understood in the cellular dynamic context, since it ultimately affects the whole metabolism. New systems biology approaches will advance our knowledge in obtaining global insights into the microbial metabolism of HMW PAHs and how the bacterial genome is operated under given environmental conditions. Although genome sequences are being analyzed, most of their functions have been predicted based on homology to other enzymes and they often contain annotations based on theoretical modeling or incomplete evidence. Therefore, functions of the numerous individual genes and their regulatory mechanisms still need to be studied at the genetic level. We still need genetic techniques such as gene cloning and gene functional analysis performed by mutational analysis. Chemical and biochemical evidence is also necessary to elucidate complete metabolic pathways since there are still many metabolite gaps in PAH degradation. All of these approaches would complement each other, which is essential for a better understanding of the multienzymatic steps in HMW PAH degradation.

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Genomic View of Mycobacterial HMW PAH Degradation Johnsen AR, Wick LY, Harms H (2005) Principles of microbial PAH-degradation in soil. Environ Pollut 133: 71–84. Juhasz AL, Naidu R (2000) Bioremediation of high molecular weight polycyclic aromatic hydrocarbons: a review of the microbial degradation of benzo[a]pyrene. Int Biodet Biodeg 45: 57–88. Kanaly RA, Harayama S (2000) Biodegradation of highmolecular-weight polycyclic aromatic hydrocarbons by bacteria. J Bacteriol 182: 2059–2067. Kazunga C, Aitken MD (2000) Products from the incomplete metabolism of pyrene by polycyclic aromatic hydrocarbon-degrading bacteria. Appl Environ Microbiol 66: 1917–1922. Kelley I, Cerniglia CE (1995) Degradation of a mixture of high-molecular-weight polycyclic aromatic hydrocarbons by a Mycobacterium strain, PYR-1. J Soil Contam 4: 77–91. Kelley I, Freeman JP, Cerniglia CE (1990) Identification of metabolites from degradation of naphthalene by a Mycobacterium sp. Biodegradation 1: 283–290. Khan AA, Wang RF, Cao WW, Doerge DR, Wennerstrom D, Cerniglia CE (2001) Molecular cloning, nucleotide sequence, and expression of genes encoding a polycyclic aromatic ring dioxygenase from Mycobacterium sp. strain PYR-1. Appl Environ Microbiol 67: 3577–3585. Kim SJ, Kweon O, Freeman JP, Jones RC, Adjei MD, Jhoo JW, Edmondson RD, Cerniglia CE (2006) Molecular cloning and expression of genes encoding a novel dioxygenase involved in low- and highmolecular-weight polycyclic aromatic hydrocarbon degradation in Mycobacterium vanbaalenii PYR-1. Appl Environ Microbiol 72: 1045–1054. Kim SJ, Kweon O, Jones RC, Edmondson RD, Cerniglia CE (2008) Genomic analysis of polycyclic aromatic hydrocarbon degradation in Mycobacterium vanbaalenii PYR-1. Biodegradation 19: 859–881. Kim SJ, Kweon O, Jones RC, Freeman JP, Edmondson RD, Cerniglia CE (2007) Complete and integrated pyrene degradation pathway in Mycobacterium vanbaalenii PYR-1 based on systems biology. J Bacteriol 189: 464–472. Kim YH, Engesser KH, Cerniglia CE (2003) Two polycyclic aromatic hydrocarbon o-quinone reductases from a pyrene-degrading Mycobacterium. Arch Biochem Biophys 416: 209–217. Kim YH, Freeman JP, Moody JD, Engesser KH, Cerniglia CE (2005) Effects of pH on the degradation of phenanthrene and pyrene by Mycobacterium vanbaalenii PYR-1. Appl Microbiol Biotechnol 67: 275–285. Kim YH, Moody JD, Freeman JP, Engesser KH, Cerniglia CE (2004) Evidence for the existence of PAHquinone reductase and catechol-O-methyltransferase

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in Mycobacterium vanbaalenii PYR-1. J Ind Microbiol Biotechnol 31: 507–516. Krivobok S, Kuony S, Meyer C, Louwagie M, Willison JC, Jouanneau Y (2003) Identification of pyreneinduced proteins in Mycobacterium sp. strain 6PY1: evidence for two ring-hydroxylating dioxygenases. J Bacteriol 185: 3828–3841. Kweon O, Kim SJ, Baek S, Chae JC, Adjei MD, Baek DH, Kim YC, Cerniglia CE (2008) A new classification system for bacterial Rieske non-heme iron aromatic ring-hydroxylating oxygenases. BMC Biochem 9: 11. Kweon O, Kim SJ, Jones RC, Freeman JP, Adjei MD, Edmondson RD, Cerniglia CE (2007) A polyomic approach to elucidate the fluoranthene degradative pathway in Mycobacterium vanbaalenii PYR-1. J Bacteriol 189: 4635–4647. Leys NM, Ryngaert A, Bastiaens L, Wattiau P, Top EM, Verstraete W, Springael D (2005) Occurrence and community composition of fast-growing Mycobacterium in soils contaminated with polycyclic aromatic hydrocarbons. FEMS Microbiol Ecol 51: 375–388. Lo´pez Z, Vila J, Grifoll M (2005) Metabolism of fluoranthene by mycobacterial strains isolated by their ability to grow in fluoranthene or pyrene. J Ind Microbiol Biotechnol 32: 455–464. Miller CD, Hall K, Liang YN, Nieman K, Sorensen D, Issa B, Anderson AJ, Sims RC (2004) Isolation and characterization of polycyclic aromatic hydrocarbon-degrading Mycobacterium isolates from soil. Microb Ecol 48: 230–238. Moody JD, Freeman JP, Fu PP, Cerniglia CE (2004) Degradation of benzo[a]pyrene by Mycobacterium vanbaalenii PYR-1. Appl Environ Microbiol 70: 340–345. Pagnout C, Frache G, Poupin P, Maunit B, Muller JF, Ferard JF (2007) Isolation and characterization of a gene cluster involved in PAH degradation in Mycobacterium sp. strain SNP11: expression in Mycobacterium smegmatis mc2155. Res Microbiol 158: 175–186. Penning TM, Burczynski ME, Hung CF, McCoull KD, Palackal NT, Tsuruda LS (1999) Dihydrodiol dehydrogenases and polycyclic aromatic hydrocarbon activation: generation of reactive and redox active o-quinones. Chem Res Toxicol 12: 1–18. Pinyakong O, Habe H, Omori T (2003) The unique aromatic catabolic genes in sphingomonads degrading polycyclic aromatic hydrocarbons (PAHs). J Gen Appl Microbiol 49: 1–19. Ramos JL, Marques S, Timmis KN (1997) Transcriptional control of the Pseudomonas TOL plasmid catabolic operons is achieved through an interplay of host factors and plasmid-encoded regulators. Annu Rev Microbiol 51: 341–373.

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26 Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs M. Seeger1,* . D. H. Pieper2 1 Laboratorio de Microbiologı´a Molecular y Biotecnologı´a Ambiental, Millennium Nucleus of Microbial Ecology and Environmental Microbiology and Biotechnology, Departamento de Quı´mica, Universidad Te´cnica Federico Santa Marı´a, Avenida Espan˜a, Valparaı´so, Chile *[email protected] 2 Biodegradation Research Group, Division of Microbiology, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1180

2 2.1 2.1.1 2.1.2 2.1.3 2.1.4 2.1.5 2.1.6

Aerobic Metabolism of PCBs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1181 Upper Pathway Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1181 Biphenyl 2,3-Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1183 cis-2,3-Dihydro-2,3-Dihydroxybiphenyl Dehydrogenases . . . . . . . . . . . . . . . . . . . . . . . . . 1183 2,3-Dihydroxybiphenyl 1,2-Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1183 2-Hydroxy-6-Phenyl-6-Oxohexa-2,4-Dienoate (HOPDA) Hydrolases . . . . . . . . . . . 1185 BphK Glutathione-S-Transferase . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1185 Lower Pathways for the Degradation of 2-Hydroxypenta-2,4-Dienoates and Benzoates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1186 2.2 Archetype bph Gene Clusters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1187 2.3 Genome Analyses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1187 2.4 Toxicity of PCBs and Their Metabolites and Bacterial Stress Response . . . . . . . . . . 1189 2.5 Metabolic Versatility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1190 2.5.1 Diversity of Rieske Non-Heme Iron Oxygenases Involved in Biphenyl Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1190 2.5.2 Mosaic Routes for Biphenyl Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1191 2.6 Optimized Enzymes and PCB Degrading Organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1193 3

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1194

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_82, # Springer-Verlag Berlin Heidelberg, 2010

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26

Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

Abstract: Microbial metabolism is responsible for the removal of persistent organic pollutants including polychlorinated biphenyls (PCBs) from the environment. Anaerobic dehalogenation processes of highly and moderately chlorinated biphenyls generate moderately and low chlorinated congeners, which can be subject to aerobic degradation, performed by enzymes of the biphenyl (bph) upper and lower pathways. These enzymes and their substrate specificities are discussed in Section 2.1. Biphenyl 2,3-dioxgenases are generally considered key enzymes of this pathway, which determine substrate range, and extent of PCB-degradation, however, also the specificity of subsequent enzymes is crucial for productive metabolism. PCB metabolism is described in detail for some model organisms, and the genetic organization of gene clusters of model organisms is described in Section 2.2. So far, the genomes of two important PCB metabolizing model organisms, namely Burkholderia xenovorans LB400 and Rhodococcus jostii RHA1 have been sequenced with the rational to better understand their overall physiology and evolution as described in Section 2.3. This has also allowed a better evaluation into genome and proteome-wide defenses against PCB toxicity, which is summarized in Section 2.4. However, more and more studies also indicate that our knowledge on enzymes and genes involved in PCB metabolism is still rather fragmentary and on overview of the diversity of enzymes reported and mosaic routes is given on Section 2.5. Finally, strategies to optimize microorganisms for improved PCB degradation and bioremediation are discussed in Section 2.6.

1

Introduction

Sustainable development aims to meet human needs while preserving the environment for future generations. The industrialization of many regions of the world has increased the environmental pollution. The removal of pollutants from the environment and the recovery of contaminated sites are major challenges of the XXI century. The Stockholm Convention of 2001 promotes the worldwide reduction and elimination of the emission of persistent organic pollutants (POPs) into the environment. Polychlorobiphenyls (PCBs), which are widely distributed in the environment, mainly in aquatic and soil ecosystems (Gomez-Gutie´rrez et al., 2007; Palma-Fleming et al., 2008) were classified in the list of the twelve POPs for priority action. Biphenyl is an aromatic compound of two bound benzene rings, which occurs naturally in coal tar, crude oil, and natural gas. The industrial chlorination of biphenyl produces a mixture of PCBs carrying 1–10 chlorine atoms. There are 209 PCB congeners that differ in position and number of the chlorines. Industrial applications of PCBs started in 1929 in U.S.A. by Monsanto. These compounds were used mainly as dielectric fluids in capacitors and transformers, but also as flame-retardants, plasticizers and ink solvents. Commercial mixtures typically consisting of 40–70 congeners were sold under trade names as Askarel and Aroclor (Monsanto, U.S.A., Canada and United Kingdom), Clophen (Bayer, Germany), Kanechlor (Kanegafuchi, Japan), Phenoclor (Prodelec, France and Spain), and Sovol and Sovtol (Orgsteklo, Orgsintez, former Soviet Union). More than 1.7 million tons of PCBs were produced worldwide (Stockholm Convention), and an important amount of these compounds have been released into the environment (Pieper and Seeger, 2008). Although adverse health effects were first recorded in the 1930s (Drinker et al., 1937), PCBs continued to be used for decades. Since then, PCBs have been shown to cause cancer (Mayes et al., 1998) and a number of serious effects on the immune, reproductive, nervous and endocrine system (Faroon et al., 2001). Some coplanar PCBs have dioxin-like properties, and

Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

26

are among the most toxic congeners. The toxicity, carcinogenicity, persistence and tendency of PCBs to bioaccumulate are of increasing environmental and health concern in many countries (Pieper and Seeger, 2008). Technologies for the treatment of PCB contaminated sites such as incineration, landfilling, thermal desorption, and chemical dehalogenation can be costly and usually involve dredging or excavation followed by disposal (Caruana, 1997). Microorganisms play a main role in the carbon cycle and in the removal of persistent organic pollutants from the environment. Bioremediation has been applied successfully for the removal of petroleum contamination. For clean-up of PCB-contaminated environments, bioremediation is a promising technology (Pieper and Seeger, 2008). Despite their chemical stability, diverse microbes have been reported as being capable to deal with PCBs and anaerobic consortia of microorganisms as well as aerobic bacteria biotransform or even mineralize PCBs. Generally, highly and moderately chlorinated PCBs are susceptible to a process termed reductive dehalogenation, in which PCBs are used as an alternative terminal electron acceptor in anaerobic respiration. The reductive dehalogenation of PCBs is congener-specific, and generally, involves selective dechlorination from para and meta positions, while chlorines at ortho position are preserved. However, ortho dechlorination of PCBs has also been reported. The first organisms capable to carry out such dehalogenations are available and belong to either the genus Dehalococcoides or Dehalobium (Cutter et al., 2001; Fennell et al., 2004; Wu et al., 2002). Even though various reductive dehalogenases for dehalogenation of tetrachloroethene, vinylchloride or chlorobenzene (Adrian et al., 2007; Mu¨ller et al., 2004; Neumann et al., 1996) have been described, enzymes involved in reductive dehalogenation of PCBs remain to be identified. Lower, but also some moderately chlorinated PCBs are susceptible to aerobic metabolism as described below.

2

Aerobic Metabolism of PCBs

Since the pioneering studies of Lunt and Evans (1970), diverse aerobic bacteria belonging to genera such as Pseudomonas, Burkholderia, Comamonas, Cupriavidus, Sphingomonas, Acidovorax, Rhodococcus and Bacillus capable of using biphenyl as a sole source of carbon and energy and capable to oxidize PCBs have been described (Pieper and Seeger, 2008).

2.1

Upper Pathway Enzymes

Based on the analysis of various biphenyl degrading isolates it could be deduced that, in general, lower chlorinated congeners are more easily transformed compared to higher chlorinated congeners and that PCB congeners with chlorines on one aromatic ring are more easily degraded than those bearing chlorine substituents on both aromatic rings. However, each isolate exhibits a particular activity spectrum with regard to the type and extent of PCB congeners metabolized, with some strains having a narrow spectrum and others, notably B. xenovorans LB400 being able to transform a broad range of congeners (Bopp, 1986; Seeger et al., 1995a, b). The degradation of biphenyl and transformation of PCBs is usually catalyzed by enzymes encoded by the so-called biphenyl (bph) upper and lower pathways (> Fig. 1).

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Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

. Figure 1 Pathways for biphenyl degradation.

1182

Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

2.1.1

26

Biphenyl 2,3-Dioxygenases

Like the degradation of various other aromatics, the degradation of biphenyl is initiated by Rieske non-heme iron oxygenases, multicomponent enzyme complexes composed of a terminal oxygenase component (iron-sulfur protein [ISP]) and different electron transport proteins (a ferredoxin and a reductase or a combined ferredoxin-NADH-reductase) (Gibson and Parales, 2000) (See > Chapter 4, Vol. 2, Part 2; > Chapter 18, Vol. 2, Part 4). Biphenyl 2,3dioxygenases (BphA) usually belong to the toluene/biphenyl branch of Rieske non-heme iron oxygenases (Gibson and Parales, 2000) where a ferredoxin (BphA3) and a ferredoxin reductase (BphA4) act as an electron transport system to transfer electrons from NADH to the terminal oxygenase, which consists of two subunits (BphA1A2), with the a-subunit being the major determinant of substrate specificity. The biphenyl 2,3-dioxygenases play a crucial role for the PCB-degradation spectra. On one side, their regiospecificity of dioxygenation of the substrate determines the sites of attack by the subsequent enzymes of the pathway while, on the other side, their substrates crucially determine the spectrum of PCB congeners that can be transformed by an organism. Studies on various biphenyl 2,3-dioxygenases have revealed considerable differences in their congener selectivity patterns, as well as their preference of the attacked ring (McKay et al., 1997; Seeger et al., 1999). The biphenyl pathway of strain LB400 oxidizes an unusually wide range of PCB (from monochlorobiphenyls to 2,3,4,5,20 ,50 -hexachlorobiphenyl, Seeger et al., 1999). Most primary catabolites that are dioxygenated by BphA of strain LB400 at ortho and meta carbons (> Fig. 2) are further metabolized by the other enzymes of the upper pathway. In contrast, dioxygenation at meta and para positions results in channeling into a dead-end pathway. Dehalogenation by BphA of ortho-chlorinated (> Fig. 2), -brominated and -fluorinated biphenyls has been observed (Haddock et al., 1995; Seeger et al., 1995a), in addition to denitration and dehydroxylation (Seeger et al., 2001). Noteworthy, the dihydroxylation of natural and synthetic isoflavonoids by BphA of strain LB400 has also been described (Seeger et al., 2003).

2.1.2

cis-2,3-Dihydro-2,3-Dihydroxybiphenyl Dehydrogenases

The second step in the metabolic pathway, the dehydrogenation of (chlorinated) cis-2,3-dihydro2,3-dihydroxybiphenyls (biphenyl 2,3-dihydrodiol) to give (chlorinated) 2,3-dihydroxybiphenyl, is catalyzed by cis-2,3-dihydro-2,3-dihydroxybiphenyl dehydrogenases (BphB, > Fig. 1). Cis-dihydrodiol dehydrogenases are involved in various aromatic degradation pathways. They are usually members of the family of short-chain alcohol dehydrogenases, generally of broad substrate specificity and able to transform several cis-dihydrodiol substrates (Jouanneau and Meyer, 2006; Rogers and Gibson, 1977). The cis-2,3-dihydro-2,3-dihydroxybiphenyl dehydrogenase of strain LB400 is able to rearomatize isoflavonoids dihydroxylated by BphA and the resulting products are assumed to have improved antioxidant properties (Arora et al., 1998).

2.1.3

2,3-Dihydroxybiphenyl 1,2-Dioxygenases

The ring-cleavage of dihydroxylated aromatic intermediates can be catalyzed by enzymes from one of two structurally and mechanistically distinct enzyme classes (See > Chapter 4, Vol. 2, Part 2). While intradiol dioxygenases, which cleave the aromatic nucleus between the hydroxyl

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Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

. Figure 2 Transformation of 4,40 -dichloro-, 2,20 -dichloro-, and 2,5,20 -trichlorobiphenyl by biphenyl 2,3-dioxygenase of B. xenovorans LB400. Unstable intermediates are shown in brackets. 4,40 -Dichlorobiphenyl is exclusively subject to 2,3-dioxygenation yielding a 2,3-dihydrodiol as product. 2,20 -Dichlorobiphenyl is dioxygenated such that one of the vic-hydroxyl groups in the cis-dihydrodiol is bound to the same carbon as the chloro-substituent. From such an unstable vic-dihydrodiol, the chloro-substituent is spontaneously eliminated. 2,5,20 -Trichlorobiphenyl is subject to both 20 ,30 -dioxygenation as well as 3,4-dioxygenation (Haddock et al., 1995; Seeger et al., 1995a, 1999, 2001).

Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

26

substituents (ortho-cleavage) use non-heme Fe(III), extradiol dioxygenases, which cleave the aromatic nucleus adjacent to the hydroxyl substituents (meta-cleavage) typically use non-heme Fe(II) for cleavage (Harayama and Rekik, 1989) even though Mn(II) dependent extradiol dioxygenases have also been reported (Hatta et al., 2003). Among the extradiol dioxygenases, three types of enzymes could be identified (See > Chapter 4, Vol. 2, Part 2). Type I extradiol dioxygenases belong to the vicinal oxygen chelate superfamily (Gerlt and Babbitt, 2001), type II enzymes are exemplified by protocatechuate 4,5-dioxygenases and are often composed of two different subunits and type III enzymes (such as gentisate 1,2-dioxygenase) belong to the cupin superfamily (Dunwell et al., 2001). Even though belonging to different families, all three types of extradiol dioxygenases share similar active sites and all type I, type II and various type III enzymes have the same iron ligands, two histidine and one glutamate, that constitute the 2-His 1-carboxylate structural motif. 2,3-Dihydroxybiphenyl 1,2-dioxygenases (BphC) involved in biphenyl degradation, usually belong to the subfamily 3A of type I extradiol dioxygenases (Eltis and Bolin, 1996) and are specialized for transformation of 2,3-dihydroxybiphenyls (> Fig. 1). Even though BphC enzymes differ in substrate specificity, they seem to be generally capable of transforming various chlorosubstituted derivatives (Dai et al., 2002; McKay et al., 2003). However, both 3,4-dihydroxybiphenyl as well as 20 -chlorosubstituted 2,3-dihydroxybiphenyls strongly inhibit BphC enzymes (Lloyd-Jones et al., 1995; McKay et al., 2003). A special feature of extradiol dioxygenases is their susceptibility to inactivation due to a rapid oxidation of the active site ferrous iron into its ferric form with concomitant loss of activity (Vaillancourt et al., 2002). Specifically 20 -chlorosubstituted 2,3-dihydroxybiphenyls promote such inactivation and thus interfere with the degradation of other compounds (Dai et al., 2002). However, significant differences between different isoenzymes were observed (Fortin et al., 2005).

2.1.4

2-Hydroxy-6-Phenyl-6-Oxohexa-2,4-Dienoate (HOPDA) Hydrolases

The fourth step in the bph pathway is catalyzed by 2-hydroxy-6-phenyl-6-oxohexa-2,4dienoate (HOPDA) hydrolase BphD, which hydrolyzes HOPDA to 2-hydroxypenta-2,4-dienoate and benzoate (> Fig. 1). HOPDA hydrolases belong to the family of C-C hydrolase enzymes of the a/b-hydrolase enzyme superfamily (Ollis et al., 1992). Studies on B. xenovorans LB400 and Rhodococcus globerulus P6 BphDs have revealed that this enzyme may be a bottleneck for the metabolism of certain PCB congeners (Seah et al., 2000, 2001; Seeger et al., 1995b). Although some differences in turnover were observed, both enzymes were similar in that HOPDAs bearing chlorine substituents at the phenyl moiety were efficiently transformed, whereas HOPDAs bearing chlorine substituents on the dienoate moiety were poor substrates and competitively inhibit BphD (see > Fig. 3). Recent studies suggest that this inhibition is due to inhibition of the histidine-mediated enol-keto tautomerization which precedes hydrolysis by BphD (Bhowmik et al., 2007).

2.1.5

BphK Glutathione-S-Transferase

BphK is a glutathione S-transferase (GST) that occurs in some bph pathways (Bartels et al., 1999). BphK was shown not to be essential for degradation of biphenyl (Bartels et al., 1999),

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Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

. Figure 3 Transformation of chlorosubstituted 2-hydroxy-6-oxo-6-phenyl-2,4-dienoates (HOPDAs) by BphD and BphK gene products, examplified by the metabolism of 4,40 -dichlorobiphenyl (Fortin et al., 2006; Gilmartin et al., 2003; Seah et al., 2000).

however, this enzyme can catalyze dehalogenation of 4-chlorobenzoate (> Fig. 3), the product of 4-chlorobiphenyl degradation by the enzymes BphA, BphB, BphC and BphD (Gilmartin et al., 2003) suggesting that BphK was recruited to facilitate the degradation of PCBs. However, 3-chloro-2-hydroxy-6-oxo-6-phenyl-2,4-dienoates, compounds that are produced by the cometabolism of PCBs by BphA, BphB and BphC (Fortin et al., 2006) and that inhibit BphD (> Fig. 3) were significantly better substrates for the enzyme compared to 4-chlorobenzoate, and were rapidly dehalogenated. Thus, BphK probably contributes to superior PCB metabolizing activities by decreasing the inhibition of BphDs by chlorinated HOPDAs.

2.1.6

Lower Pathways for the Degradation of 2-Hydroxypenta-2,4Dienoates and Benzoates

The metabolism of (chloro)biphenyls by the biphenyl upper pathway results, in the best case, in the formation of (chlorinated) 2-hydroxypenta-2,4-dienoates and (chlorinated) benzoates (> Fig. 1). 2-Hydroxypenta-2,4-dienoate is transformed by 2-hydroxypenta-2,4-dienoate hydratase (bphH), 4-hydroxy-2-oxovalerate aldolase (bphI) and an acylating acetaldehyde dehydrogenase (bphJ) to pyruvate and acetyl-CoA (> Fig. 1), which then can enter the Krebs cycle. Thus, these enzymes should allow growth of bacterial strains on biphenyls chlorinated at one aromatic ring only, which yield chlorinated benzoates as dead-end metabolites and unchlorinated 2-hydroxypenta-2,4-dienoate. If chlorinated 2-hydroxypenta-2,4dienoate can be transformed has yet to be elucidated. Besides 2-hydroxypenta-2,4-dienoates, benzoates are generated during BphD catalyzed hydrolysis of HOPDAs (> Figs. 1 and > 3). Benzoate is a growth substrate for a broad range of Actinobacteria and Proteobacteria and under aerobic conditions can be mineralized either via catechol and a 3-oxoadipate pathway or via 2,3-dihydrodihydroxybenzoyl-CoA and nonoxygenolytic cleavage of the aromatic ring (See > Chapter 4, Vol. 2, Part 2). In contrast, chlorobenzoates, typically formed during metabolism of PCBs by the biphenyl upper pathway, are usually dead-end metabolites for PCB transforming bacteria (for bacterial strategies to degrade chloroaromatics including chlorobenzoates, See > Chapter 5, Vol. 2, Part 2).

Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

2.2

26

Archetype bph Gene Clusters

Our knowledge on biphenyl degradation and PCB metabolism is significantly governed by analysis of some isolates which have been described in detail and are regarded as the archetype PCB degraders, among them, strains B. xenovorans LB400 and R. jostii RHA1, whose genomes have recently been deciphered. B. xenovorans LB400 (Mondello, 1989), P. pseudoalcaligenes KF707 (Furukawa and Miyazaki, 1986) and others harbor an operon comprising genes encoding enzymes of the biphenyl upper pathway, a glutathione S-transferase (bphK) and genes encoding enzymes involved in the transformation of 2-hydroxypenta-2,4-dienoate released during hydrolysis of HOPDA (> Fig. 4). Regulation of these clusters is assumed to be mediated by an orf0 encoded GntR family transcriptional regulator (Beltrametti et al., 2001; Watanabe et al., 2000). P. putida KF715 contains a bphABCD gene cluster (Hayase et al., 1990) (> Fig. 4) which was suggested to have evolved from a LB400-type gene cluster. In LB400, the bph genes are located on a genomic island on the mega plasmid (Chain et al., 2006). The presence of bph genes on mobile genetic elements indicate that these genes are able to move between genomes, thus allowing adaptation of microbial communities to PCBs. A second type of bph gene cluster was observed in Acidovorax sp. strain KKS102 (Kikuchi et al., 1994) and Cupriavidus oxalaticus A5 (Springael et al., 1993) (bphSEGF(orf4) A1A2A3A4BCD(orf1)A4) (> Fig. 4). In these clusters, genes encoding enzymes involved in the transformation of 2-hydroxypenta-2,4-dienoate (designated bphEGF) are preceding genes encoding upper pathway enzymes, and the gene encoding the reductase subunit of biphenyl dioxygenase (bphA4) is localized at the end of the gene cluster. Like in LB400, regulation was shown to be dependent on a member of the GntR family of transcriptional regulators (BphS) (Mouz et al., 1999) and at least in C. oxalaticus A5 the bph are also located on a mobile genomic island (Toussaint et al., 2003). A bphAaAbAcAdCB gene cluster devoid of a gene encoding a HOPDA hydrolase was observed in R. jostii RHA1 (Masai et al., 1995) (> Fig. 4) localized on the linear plasmid pRHL1 (Takeda et al., 2004). This catabolic gene cluster, like the similarly structured bph gene clusters of Rhodococcus sp. M5 (Peloquin and Greer, 1993) is regulated by a two-component signal transduction system composed of a BphT response regulator and a BpdS sensor kinase, promoting transcriptional induction by a variety of aromatic compounds (Takeda et al., 2004). A nearly identical plasmid localized gene cluster has been shown to be involved in isopropylbenzene degradation by R. erythropolis BD2 (Stecker et al., 2003), indicating such gene clusters to be involved in the degradation of differently substituted aromatics.

2.3

Genome Analyses

So far, the genomes of two potent PCB-degrading bacteria, B. xenovorans LB400 (Chain et al., 2006) and R. jostii RHA1 (McLeod et al., 2006) have been sequenced with the rational to better understand their overall physiology and to foster their applicability for bioremediation purposes. The LB400 genome has a size of 9.73 Mbp distributed over two circular chromosomes (4.87 and 3.36 Mbp, respectively) and a circular megaplasmid (1.47 Mbp). Strain RHA1 has a genome of 9.70 Mbp arranged on a linear chromosome (7.80 Mbp) and three linear plasmids (1.12, 0.44 and 0.33 Mbp, respectively). Both strains inhabit soil and plant rhizosphere niches. The large genomes of strains LB400 and RHA1 have evolved by different means. More than 20% of the genome of strain LB400 was recently acquired via horizontal gene transfer (HGT).

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. Figure 4 Genetic organization of the bph gene clusters of B. xenovorans LB400, P. putida KF715, Rhodococcus sp. strain M5, Acidovorax sp. strain KKS102, Rhodococcus sp. K37, Bacillus sp. JF8, Sphingobium yanoikuyae B1, and of the bph and etb gene clusters of R. jostii RHA1.

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26

In contrast, strain RHA1 evolved through ancient acquisition or gene duplication and acquired far fewer genes by recent HGT than LB400 (McLeod et al., 2006). Both bacterial strains have an unusually high metabolic versatility for degradation of aromatic compounds both with respect to peripheral routes activating aromatics for ringcleavage as well as for central routes channeling those intermediates into the Krebs cycle (See also > Chapter 4, Vol. 2, Part 2). The genes encoding enzymes of the biphenyl upper bph pathway are located in both strains on acquired and mobile genetic elements. In LB400 they are encoded by a genomic island on the megaplasmid, indicating that these genes were acquired via HGT (Chain et al., 2006). The genomic islands also provide strain LB400 with other highly specialized metabolic capabilities such as the abilities to degrade 2-aminophenol or 3-chlorocatechol. In strain RHA1, the bph genes are, like 11 of the 26 peripheral aromatic pathways, located on the plasmids (McLeod et al., 2006).

2.4

Toxicity of PCBs and Their Metabolites and Bacterial Stress Response

The toxicity of POPs and their catabolites for microorganisms is a major challenge for bioremediation processes (Blasco et al., 1995; Camara et al., 2004). PCBs are expected to accumulate in bacterial membranes due to their lipophilic character (Sikkema et al., 1995) (See also > Chapter 52, Vol. 2, Part 9) and, in fact, PCBs decreases bacterial cell viability (Ca´mara et al., 2004). Noteworthy, some metabolic intermediates are even more toxic than PCBs. Degradation of specific PCB congeners by diverse bacteria is incomplete with a concomitant accumulation of different metabolic intermediates (Seah et al., 2000; Seeger et al., 1995a). Biotransformation of PCBs by BphA and BphB produces dihydrodiols and dihydroxybiphenyls, which are highly toxic for bacteria (Ca´mara et al., 2004). The increased polarity of dihydroxylated metabolites increases their aqueous solubility, contributing to this toxic effect. Hydroxylated PCB metabolites can affect the DNA content of bacteria, inhibiting bacterial cell separation (Hiraoka et al., 2002). The conversion of PCBs compounds into products with increased toxicity is also known from the bioactivation of xenobiotics and drugs in higher organisms. In fact the oxidation by cytochrome P450 generate reactive products that can be cytotoxic (> Fig. 5). Moreover, chlorobenzoates, which are often dead-end products by PCB metabolizing bacteria, can be transformed into deleterious downstream products. 3-Chlorocatechol can inactivate extradiol dioxygenases such as 2,3-dihydroxybiphenyl 1,2-dioxygenases (Vaillancourt et al., 2002), thus interfering with the biphenyl upper pathway. Channeling of 4-chlorocatechol into the wide-spread 3-oxoadipate pathway can result in formation of the antibiotic protoanemonin (Blasco et al., 1995), and protoanemonin was assumed to be the reason for the poor survival of PCB cometabolizing organisms in soil microcosm studies (Blasco et al., 1997). Toxicity of PCBs a direct result of the production of deleterious metabolites during cometabolism was also indicated in studies using the PCB degraders LB400 and RHA1 (Parnell et al., 2006). Although PCBs were shown to partition to the cell fraction, no significant effects were observed regarding viability or growth rate in either strain under non PCB-degrading conditions whereas significant strain dependent differences were observed in cells metabolizing PCBs. Strain LB400 exhibited a high tolerance to PCB degradation-dependent toxicity whereas RHA1 was highly sensitive.

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Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

. Figure 5 Biotransformation of PCBs into cytotoxic metabolic intermediates. Cytotoxic metabolites are boxed. Inhibition is indicated by a dashed arrow.

Evaluation of the genome and proteome-wide defenses against PCB toxicity in LB400 showed induction of the molecular chaperones DnaK and GroEL during (chloro)biphenyl degradation (Agullo´ et al., 2007) and of DnaK and HtpG by 4-chlorobenzoate, a dead-end metabolite of the biphenyl upper pathway (Martı´nez et al., 2007), indicating that such exposure constitutes stressful conditions. Oxidative stress through the generation of reactive oxygen species (Chavez et al., 2004), probably resulting from the action of oxygenases in the metabolism, resulted in the induction of the alkyl hydroperoxide reductase AhpC (Agullo´ et al., 2007), which detoxifies peroxides and induction of a putative chloroacetaldehyde dehydrogenase (Denef et al., 2005) was suggested to reduce the concentration of toxic chlorinated aliphatic compounds resulting from PCB degradation. In order to establish optimized bioremediation processes for PCBs, it will be of paramount importance to overcome dead-end steps in the catabolic process and to balance the activities of enzymes involved in the degradation to avoid accumulation of toxic metabolites.

2.5

Metabolic Versatility

2.5.1

Diversity of Rieske Non-Heme Iron Oxygenases Involved in Biphenyl Metabolism

More and more information becomes currently available that Rieske type non-heme iron oxygenases outside of the archetype toluene/biphenyl branch are involved in biphenyl degradation. As an example, the bph operon of Bacillus sp. JF8 harbors a bphRDA1A2BC cluster

Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

26

(Mukerjee-Dhar et al., 2005) (> Fig. 4) encoding enzymes only distantly related to enzymes of archetype Bph enzymes (> Fig. 6) and BphA1 is more closely related to naphthalene dioxygenases NidA from Rhodococcus sp. strain I24 (Larkin et al., 1999). Also, the Mn(II) dependent BphC and BphD evidently belong to new subfamilies in the phylogeny of extradiol dioxygenases and hydrolases acting on extradiol cleavage products (Hatta et al., 2003; MukerjeeDhar et al., 2005). Analysis in Sphingobium yanoikuyae strain B1 revealed that a single ferredoxin and a single ferredoxin reductase, encoded by bphA3 and bphA4, respectively, can be shared by multiple oxygenase systems (Bae and Kim, 2000), including biphenyl oxygenase encoded by the bphA1fA2f genes (Yu et al., 2007). In a phylogenetic analysis, BphA1f does not cluster with known BphAs, but is more related with PhnI from Sphingomonas sp. strain CHY-1, which was shown to be able to oxidize at least 8 PAHs made of 2–5 aromatic rings (Demaneche et al., 2004; Jakoncic et al., 2007) (> Fig. 6). Accordingly, BphA1f is responsible for the capability of S. yanoikuyae B1 to dihydroxylate large aromatic compounds, such as chrysene and benzo[a] pyrene (Ferraro et al., 2007). Also the etbA1 encoded oxygenase a-subunit of R. jostii RHA1, only distantly related to previously characterized BphA1 proteins (see > Fig. 6), has been implicated to be important for PCB metabolism as it is more active on highly chlorinated congeners than the bphAa encoded one (Iwasaki et al., 2006) and obviously appropriate for both biphenyl and ethylbenzene transformation. Furthermore, another type of biphenyl oxygenase a-subunit has been discovered in Rhodococcus sp. strain K37 (Taguchi et al., 2007) (> Fig. 6), evidencing that diversity of oxygenases involved in biphenyl degradation is highly underestimated. In general, it has to be considered that, despite the evolutionary adaptation of enzymes for specific substrates, the enzymes of a particular pathway often catalyze the transformation of a range of substrate analogues and specifically Rieske non-heme iron oxygenases are described by a broad substrate specificity. Among various other oxygenases, chlorobenzene dioxygenases (belonging, like biphenyl dioxygenases, to the toluene/biphenyl branch of Rieske non-heme iron oxygenases) (Raschke et al., 2001), naphthalene dioxygenases (belonging to the naphthalene family of Rieske non-heme iron oxygenases) (> Fig. 6) (Barriault and Sylvestre, 1999), phenanthrene dioxygenases (Kasai et al., 2003) or carbazole 1,9a dioxygenases (Nojiri et al., 1999) are capable to transform biphenyl. Additionally, culture independent studies revealed the abundance of novel branches of Rieske type non-heme iron oxygenases in contaminated sites, the importance and environmental function of which still remains to be elucidated (Taylor et al., 2002; Witzig et al., 2006) and recent studies on PCB contaminated sites indicated novel undescribed types to be possibly important in situ (Leigh et al., 2007).

2.5.2

Mosaic Routes for Biphenyl Metabolism

Metabolism of biphenyl and PCBs should not be regarded as a simple linear pathway, but often necessitates the complex interplay between different catabolic gene modules even inside single strains. As an example, the bph cluster of R. jostii RHA1 does not comprise a bphD gene, and such activity has to be recruited from elsewhere in the genome. In fact, three hydrolases were shown to be upregulated during growth of RHA1 on biphenyl (Goncalves et al., 2006) with one of them, termed BphD previously shown to be capable to attack HOPDA (Yamada et al., 1998). P. putida strain CE2010 mineralizes biphenyl by a mosaic of tod (toluene) and cmt

1191

26

. Figure 6 Dendrogram showing the relatedness of oxygenase a-subunits of Rieske non-heme iron oxygenases. a-Subunits of supposed or validated biphenyl 2,3-dioxygenases are indicated by a filled circle.

1192 Genetics of Biphenyl Biodegradation and Co-Metabolism of PCBs

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(cumate) pathways (Ohta et al., 2001). As previously reported, toluene dioxygenase (TodC1C2BA), toluene dihydrodiol dehydrogenase (TodD) and the meta-cleavage enzyme TodE have a significant cross-reactivity with biphenyl or metabolites produced during biphenyl degradation (Furukawa et al., 1993), whereas TodF 2-hydroxy-6-oxohepta-2,4-dienoate hydrolase cannot cope with HOPDA. Recruitment of a hydrolase active with HOPDA, such as in RHA1 allows CE21010 to mineralize biphenyl. The same holds for extradiol dioxygenases, especially in Rhodococcus, where the presence of multiple extradiol dioxygenase encoding genes has been reported (McLeod et al., 2006; Taguchi et al., 2004). The metabolic versatility of catabolic enzymes and pathways is an indication of the ongoing evolution of bacterial metabolism, thus endowing environmental microbes with the capabilities to deal with a broad range of pollutants.

2.6

Optimized Enzymes and PCB Degrading Organisms

Pollution by PCBs typically consists of mixtures of congeners and only a fraction of these can be attacked by known BphAs. Therefore, for improved PCB catabolic pathways, recruitment or generation of improved biphenyl 2,3-dioxygenases ranges is required. The construction of chimeric BphA derivatives generated by the combination of gene segments of well known PCB-degraders enabled the identification of key domains of these oxygenases (Kimura et al., 1997; Kumamaru et al., 1998) and generated biphenyl 2,3-dioxygenases with improved capacities (Erickson and Mondello, 1993; Mondello et al., 1997; Suenaga et al., 1999, 2002). A directed evolution approach using random mutagenesis to specific segments allowed generating BphAs with increased turnover of PCBs, largely recalcitrant to attack by the parental enzyme (Zielinski et al., 2006). On the other side, the isolation of naturally occurring enzymatic activities by metagenomic methods which circumvent the cultivation of organisms has been used (Ca´mara et al., 2007). Recent studies have combined both approaches and using both, the broad natural diversity and methods of artificial evolution by family shuffling of soil DNA encoding BphA segments to generate BphA variants with novel regioselectivities (Vezina et al., 2007). Even though enzyme optimizations have been mainly applied to biphenyl 2,3-dioxygenases, efforts have been also directed towards elucidation of pathway bottlenecks in downstream enzyme activities (see Section 2.1) and in identifying optimized isoenzymes. As an example, a HOPDA hydrolase with novel specificities towards polychlorinated biphenyl metabolites, which specifically transformed 3-chlorosubstituted HOPDAs, compounds that inhibit archetype BphDs, was recently characterized from S. wittichii RW1 (Seah et al., 2007). As described above, most current available microorganisms are capable to mineralize biphenyl, but only cometabolize PCBs due to the absence of enzyme necessary for mineralization of chlorobenzoates, generated through metabolism by the biphenyl upper pathway (see chapter). The strategy of combining complementary metabolic activities for the development of microorganisms capable of mineralizing PCBs by combining an oxidative pathway for (chloro)biphenyl transformation (encoded by the bph genes) into (chloro)benzoate with a chlorobenzoate degradative pathway had been followed for various years. Several hybrid strains have been engineered by conjugative matings (Reineke, 1998) of appropriate organisms or by introduction of the bph genes into chlorobenzoate degraders, usually using a degradative pathway for chlorobenzoates via the corresponding chlorocatechols (See > Chapter 4, Vol. 2,

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Part 2). By cloning and expressing the genes encoding enzymes for ortho- and para-dechlorination of chlorobenzoates in biphenyl-degrading and chlorinated biphenyls co-metabolizing strains, derivatives capable of growing on and completely dechlorinating 2- and 4-chlorobiphenyl could also be obtained (Hrywna et al., 1999). However, it should be noted, that novel isolates with interesting metabolic properties capable to mineralize some PCBs are still being isolated (Adebusoye et al., 2008).

3

Research Needs

Significant advances have been made in recent years in the elucidation of the genetic and biochemical basis of aerobic bacterial degradation of PCBs. Detailed knowledge of the enzymatic steps and substrate specificity determinants of key enzymes as well as of different catabolic pathway segments necessary for mineralization is available. In order to optimize degradation of PCBs, bacteria with improved catabolic capabilities will be required either through novel isolation strategies or through protein engineering and recruitment of additional specific enzymes to overcome pathway bottlenecks. However, it should be noted that the recent decades have seen a large number of trial-and-error approaches to release specific bacterial strains into the environment which were mostly unsuccessful, mainly because of the multivariate nature of the systems involved, and secondly, because of an incomplete understanding of the bacterial catalytic and survival capacities under conditions of stress and the environmental factors governing those responses. The availability of complete genome sequences of PCB degraders, more of which will without doubts be available in the next years, will allow to unravel and understand full bacterial genome regulatory networks and physiology under conditions of environmental stresses and pollutant metabolism, and the ecological behavior of microorganisms in complex mixed microbial communities as present in contaminated sites. Given the facts that PCBs in the environment are present as complex congener mixtures, different catabolic key enzymes have been identified as bottlenecks for each congener, and optimization of a biocatalyst for a single congener often results in decreasing its performance on other congeners, it becomes increasingly evident that microbial communities have to become the focus of research. In contrast to single bacterial species, which are very unlikely to be able to deal with complex congener mixtures, microbial communities can be more versatile, and carbon sharing through complex metabolic interactions can increase PCB degradation. Gaining an understanding of PCB degradation will require integration of single organism studies with efforts to understand functioning of complex communities.

Acknowledgments MS gratefully acknowledges Kenneth N. Timmis for open him the door to biodegradation and for generous support. The authors would like to acknowledge financial support by grant EU ICA4-CT2002-10011. M.S. gratefully acknowledges support from the grants FONDECYT (1070507, 1020221 and 7080148), USM (130522, 130836, 130948), MILENIO P04/007-F (MIDEPLAN), PBCT RED 12 and CONICYT-BMBF. D.P. gratefully acknowledges support from the grant EU GOCE 003998 (BIOTOOL) and BACSIN.

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Wu Q, Watts JE, Sowers KR, May HD (2002) Identification of a bacterium that specifically catalyzes the reductive dechlorination of polychlorinated biphenyls with doubly flanked chlorines. Appl Environ Microbiol 68: 807–812. Yamada A, Kishi H, Sugiyama K, Hatta T, Nakamura K, Masai E, Fukuda M (1998) Two nearly identical aromatic compound hydrolase genes in a strong polychlorinated biphenyl degrader, Rhodococcus sp. strain RHA1. Appl Environ Microbiol 64: 2006–2012. Yu C, Liu W, Ferraro D, Brown E, Parales JV, Ramaswamy S, Zylstra GJ, Gibson DT, Parales RE (2007) Purification, characterization and crystallization of the components of a biphenyl dioxygenase system from Sphingobium yanoikuyae B1. J Ind Microbiol Biotechnol 34: 311–324. Zielinski M, Kahl S, Standfuss-Gabisch C, Ca´mara B, Seeger M, Hofer B (2006) Generation of novel-substrate-accepting biphenyl dioxygenases through segmental random mutagenesis and identification of residues involved in enzyme specificity. Appl Environ Microbiol 72: 2191–2199.

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27 Genetics and Molecular Features of Bacterial Dimethylsulfoniopropionate (DMSP) and Dimethylsulfide (DMS) Transformations J. M. Gonza´lez1 . A. W. B. Johnston2 . M. Vila-Costa3 . A. Buchan4 Department of Microbiology, University of La Laguna, La Laguna, Tenerife, Spain [email protected] 2 School of Biological Sciences, University of East Anglia, Norwich, UK [email protected] 3 Department of Marine Sciences, University of Georgia, Athens, Georgia, USA; and Department of Continental Ecology-Limnology, Centre d’Estudis Avanc¸ats de Blanes, CSIC, Blanes, Spain [email protected] 4 Department of Microbiology, University of Tennessee, Knoxville, Tennessee, USA [email protected]

1

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1202 2 Demethylation of DMSP: Assimilation of DMSP Sulfur into Protein . . . . . . . . . . . . . 1203 3 Genes for the Release of DMS from DMSP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1205 4 Why Make DMS in the First Place? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1206 5 Degradation of DMS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1206 6 Genes for the Transformation of DMS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1208 7 Bacteria Make the Most of DMS(P): Oxidation of Inorganic Sulfur . . . . . . . . . . . . . . 1208 8 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1209

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_83, # Springer-Verlag Berlin Heidelberg, 2010

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Genetics and Molecular Features of Bacterial DMSP and DMS Transformations

Abstract: The transformations of dimethylsulfoniopropionate (DMSP; (CH3)2S+ CH2CH2COO ) by bacterioplankton play important roles in the global sulfur cycle. This compound is produced in large quantities primarily for use as an osmolyte by marine algae. DMSP is a labile compound although the complete mineralization of DMSP is only a minor fate in the ocean. DMSP is the main precursor of dimethylsulfide (DMS; CH3-S-CH3), a radiatively active trace gas that contributes to global climate regulation. However, it is believed that the main pathway for the transformation of DMSP involves an assimilation step in which DMSP sulfur is incorporated efficiently into cell biomass, leaving relatively little sulfur available for release as DMS. DMSP is rapidly turned over in the environment and the diversity of pathways for its transformation are likely not yet fully realized. This chapter covers recent findings on the genetics of DMSP catabolism; their discoveries are changing our view of the role of this compound in the World’s oceans. Although even less is known about bacterially-mediated transformations of DMS, the handful of genes that have been described in a limited number of bacteria is also reviewed in this chapter.

1

Introduction

Dimethylsulfoniopropionate (DMSP) is a hugely abundant molecule in nature. It is produced mostly in the oceans, by ubiquitous phytoplankton, seaweeds and some species of terrestrial and aquatic vascular plants. DMSP acts primarily as an osmoprotectant for the organisms that make it, although it may also serve as an antioxidant, a cryoprotectant or a predator deterrent. Intracellular DMSP concentrations vary among organisms, and in some marine algae (e.g., dinoflagellates and prymnesiophytes) they can be as high as 0.2–0.5 M. DMSP is also an important substrate for bacteria that decompose the fraction that leaks from aged algal cells or which is liberated following viral lysis or zooplankton grazing. Once DMSP is released, it is taken up and transformed relatively quickly (Kiene and Linn, 2000) serving as both a sulfur and carbon source to bacterioplankton (Kiene et al., 2000). DMSP can support 1–15% of the marine bacterial carbon demand and all the sulfur demand (Kiene and Linn, 2000; Simo´ et al., 2002), which represent massive contributions for a single compound. Many different marine bacteria can degrade DMSP. These include strains from the alpha-, beta- and gammaproteobacteria (e.g., Roseobacter, Alcaligenes and Vibrio respectively) and several such strains can grow on DMSP as a sole carbon source. It was known early on that bacteria possess at least two different ways to catabolize DMSP. One of these involves a demethylation step, and is the predominant pathway in natural environments (Kiene et al., 1999; Simo´ and Pedro´s-Alio´, 1999). The other general mechanism for DMSP catabolism liberates the highly volatile dimethyl sulfide (DMS), a product that is of major environmental importance in its own right. Most of the DMS is recycled in the marine food web (Simo´, 2001) but 20 Tg escapes into the atmosphere each year (Kettle and Andreae, 2000) making it an important link in sulfur exchange between the oceans and the atmosphere. Furthermore, once released in the air, it is abiologically oxidized to hygroscopic compounds such as sulfuric acid and methanesulfonic acid (MSA). These attract water droplets, and act as cloud condensation nuclei to increase the albedo, regulating the radiation balance of the Earth. It was even suggested that this forms a self-regulated feedback between DMS release and the plankton communities that produce DMSP and DMS (> Fig. 1; Charlson et al., 1987).

Genetics and Molecular Features of Bacterial DMSP and DMS Transformations

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. Figure 1 Proposed main pathways for DMSP cycling in seawater and atmosphere.

Despite the importance of bacterial DMSP catabolism and DMS production, studies of the molecular genetics and detailed enzymology of these processes are in their infancy.

2

Demethylation of DMSP: Assimilation of DMSP Sulfur into Protein

The utilization of DMSP is a clear example of how bacterioplankton are adapted to optimize growth by utilizing reduced forms of a substrate (Vallino et al., 1996). Sulfate is one of the most abundant ions in natural waters, especially seawater, and its concentration is far from growth-limiting. However, the sulfate S atom is in its most oxidized state (+VI) and must be reduced to its most reduced ( II; sulfide) form during bacterial S assimilation. This major energetic cost may explain why bacteria prefer DMSP as a sulfur source, even though it is 2–3 orders of magnitude less abundant than sulfate in seawater. Not surprisingly, other sulfur substrates have been previously found to substitute for sulfate as a sulfur source, such as amino acids, which can inhibit the assimilation of sulfate (Kiene et al., 1999). Bacteria also assimilate DMSP in laboratory cultures, indicating that model organisms can be isolated as pure cultures for physiological studies. For example, Silicibacter pomeroyi

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obtained all its sulfur from DMSP in medium with a relatively high concentration of glucose (5 mM). Only when the concentrations of DMSP were high enough did a different pattern take place: S. pomeroyi released DMS into the medium (Gonza´lez et al., 1999). The widely distributed oligotrophic marine bacterium Pelagibacter ubique (a member of the SAR11 group) obtains its sulfur exclusively from DMSP or other forms of organic sulfur, since it lacks the genes necessary to assimilate S-sulfate (Tripp et al., 2008). The utilization of DMSP sulfur in S. pomeroyi and P. ubique involves the demethylation pathway, with the formation of methanethiol (MeSH), some of which is incorporated subsequently into bacterial amino acids (> Fig. 2). The enzyme cystathionine g-synthetase can incorporate either sulfide or MeSH directly into O-acyl homoserine and, in the latter case, the result is the direct formation of methionine (Kiene et al., 1999). A DMSP methyltransferase in the glycine cleavage T-family, encoded by the dmdA gene, was first identified in the Roseobacter and SAR11 clades (Howard et al., 2006). The DmdA enzyme generates methylmercaptopropionate (MMPA) plus a methyl group that enters the C1 pool via the coenzyme tetrahydrofolate (> Fig. 2). Subsequently, sulfur derived from MMPA can be incorporated into methionine via MeSH and from here to higher trophic levels.

. Figure 2 Pathways for the transformation of DMSP in seawater and genes involved. Percentages indicate the amount of carbon that goes through each pathway based on environmental data.

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No gene or enzyme that releases MeSH has been identified thus far. Alternatively, MMPA can be degraded to mercaptopropionate (3-MPA) in a second demethylation reaction and thence to acrylate and sulfide (Visscher and Taylor, 1994). In the natural environment, DMSP sulfur demethylation is a competing route for the transformation of DMSP into the climate active gas DMS (> Fig. 1). Incorporation of DMSP dominates the microbial transformation of this compound in the environment, based on bulk activity measurements (Kiene et al., 2000). Consistent with this, gene stoichiometry in metagenomic libraries shows that the dmdA homologs occur in 30% of the cells in bacterial cells in the oceans (Howard et al., 2008), whereas the ddd genes involved in pathways that generate DMS are less abundant (Curson et al., 2008; Todd et al., 2007). DMSP functions as a labile substrate that many different bacteria can exploit in the natural environment. Using microautoradiography combined with in situ oligonucleotide hybridization, the contribution of bacteria that assimilated DMSP sulfur in environmental samples was found to be similar or only slightly smaller than the fraction of bacteria that took up the amino acid leucine (Malmstrom et al., 2004; Vila et al., 2004). The seasonal pattern of uptake of both leucine and DMSP was also shown to be similar (Vila et al., 2004; Vila-Costa et al., 2007). These studies showed that taxa that are known to dominate the marine bacterioplankton communities take up DMSP. However, recent results suggest that the fate of DMSP sulfur is controlled by changes in the carbon reservoir and less by the phylogenetic composition of the bacterioplankton (Vila-Costa et al., 2007). Therefore, one would expect that enzymes and pathways for the transformation of DMSP would be widespread across different phylogenetic groups.

3

Genes for the Release of DMS from DMSP

For decades, the widely accepted route for DMSP dependent DMS production was that DMSP was cleaved by a ‘‘DMSP lyase,’’ releasing DMS, acrylate and a proton. A gene, dddL, was recently found in several marine Rhodobacterales (alphaproteobacteria) whose product has the characteristics of such a DMSP lyase (Curson et al., 2008). The DddL protein was previously in a DUF (domain of unknown function) and did not contain the N-terminal sequence that was reported for a DMSP lyase from Pseudomonas doudoroffii and of Alcaligenes (De Souza and Yoch, 1996). A second, completely different mechanism for DMS emission, and encoded by the dddD gene was found in several other bacteria (Todd et al., 2007). The dddD polypeptide is not a lyase, but may add acyl-CoA to DMSP (> Fig. 2), the resulting DMSP-CoA being predicted to liberate DMS non-enzymatically before further catabolism of the C3 compound acryloylCoA to 3-hydroxy propionate. The fact that the very different genes dddD and dddL are involved in the initial release of DMS may explain earlier findings that ‘‘DMSP lyase’’ was very different in different bacteria (Yoch, 2002). The DddL-type of DMSP lyase is confined to the Order Rhodobacterales, although not all strains, even of a single species (e.g., Rhodobacter sphaeroides) contain it (Curson et al., 2008). In contrast, the DddD acyl CoA transferase is very widespread, being found in strains of alphaand gammaproteobacteria (e.g., Sagittula and Marinomonas, respectively), and also in terrestrial bacteria that interact with roots of land plants. These include strains of the N2-fixing symbiont Rhizobium and the rhizosphere bacterium Burkholderia (Todd et al., 2007). Thus dddD may be prone to horizontal gene transfer among bacteria.

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Consistent with this, the ddd gene clusters of different bacteria that contain dddD posses other genes involved in DMSP transport (see below), catabolism or gene regulation. Strikingly, the gene products for a given function may be in wholly different gene families; thus the DMSP transporter in Marinomonas is an integral membrane-bound BCCT (betaine, choline, carnitine transport) type but Rhizobium transports DMSP by an ABC (ATP-binding cassette) system (Todd et al., 2007). In addition to these specialized DMSP transporters, most bacteria, including several known DMSP-catabolizing ones, have other versions of the BCCT-type and betaine ABC transporters that may serve for the general import of structurally similar molecules. Indeed, it was shown directly that E. coli has transporters that can import both betaine and DMSP (Cosquer et al., 1999) while Kiene et al. (1998) used bioassays with natural samples, to show that DMSP and glycine betaine share the same transporters. Also, S. pomeroyi has genes for BCCT-type and ABC-type transporters that were induced by exposure of the cells to DMSP (Bu¨rgmann et al., 2007) and a likely DMSP uptake gene, similar to opuD was noted by Moran et al. (2004).

4

Why Make DMS in the First Place?

On the face of it, the DMS-releasing pathway(s) of DMSP catabolism seem inherently wasteful, since all the sulfur and 40% of the carbon is lost as the DMS escapes. Indeed, insignificant amounts of carbon and sulfur are incorporated into cell biomass from DMS by natural assemblages (del Valle et al., 2007; Kiene et al., 1999; Vila-Costa et al., 2006). Given that bacteria can recoup all of the S and the C from DMSP via demethylation, and that some strains, such as S. pomeroyi can express both the DmdA demethylase and the DddD acyl-CoA transferase, the apparently wasteful release of DMS seems odd. Interestingly, experimental observations on S. pomeroyi grown on media with sufficient carbon (5 mM glucose) plus DMSP that served as the sole sulfur source, only liberated DMS when the concentration of DMSP was higher than 0.01 mM (Gonza´lez et al., 1999). Another Roseobacter strain, Dinoroseobacter shibae DFL 12, not only contains homologs of dmdA, and both dddA and dddL, but also has genes involved in the transformation of the products of DMSP degradation. These are a putative homolog of the DMS monooxygenase (see below) and inorganic sulfur oxidation enzymes, making it quite a versatile sulfur-transforming organism. Bacteria in the Roseobacter cluster, such as S. pomeroyi and D. shibae, make good candidates to study the physiological roles of the different types of DMSP catabolism that can be accomplished by a single strain. Additionally, it may be relevant that DMS can act as a signal molecule and that maybe at least some bacteria make DMS to either attract other organisms or in defense, perhaps together with the acrylate that is formed by the bacteria that use the DMSP lyase. Certainly, bacteria that can make DMS from DMSP do not necessarily grow on DMSP as sole carbon source, or degrade DMS any further, at least under laboratory conditions, suggesting that this process may not be wholly involved in nutrition.

5

Degradation of DMS

DMS could potentially be a source of sulfur for marine bacterioplankton since its degradation products include MeSH and sulfide (> Fig. 3). However, the DMS that is catabolized in marine

Genetics and Molecular Features of Bacterial DMSP and DMS Transformations

27

. Figure 3 Known routes for the aerobic transformation of DMS for which key genes have been identified.

systems mostly ends up as DMSO and sulfate (del Valle et al., 2007; Kiene et al., 1999; VilaCosta et al., 2006). DMS turnover rate is also lower when compared to that of DMSP (Kiene et al., 1999). The numbers of bacterial isolates that can utilize DMS as a source of carbon, sulfur or energy are not as great as those that grow on DMSP. In most cases, isolates that transform DMS have been enriched or isolated from environments, such as biofilters, with high concentrations of DMS and related compounds (Pol et al., 1994). However, bacteria that break down DMS belong to a wide range of phylogenetic groups. Hyphomicrobium spp. have been isolated after DMS enrichment (de Bont et al., 1981; Pol et al., 1994; Suylen and Kuenen, 1986). Rhodovulum sulfidophilum SH1 utilizes DMS as an electron donor during photoautotrophic growth (Hanlon et al., 1994). The Gram positive Rhodococcus sp. SY1 (Omori et al., 1995) and Arthrobacter (Borodina et al., 2000) have been demonstrated to utilize DMS. Acinetobacter sp. 20B (Horinouchi et al., 1997) grows on DMS as the sole source of sulfur, whilst Marinobacterium sp. DMS-S1 does it only in the presence of light (Fuse et al., 2000). Methanogens (Kiene et al., 1986) and sulfate-reducing bacteria (Tanimoto and Bak, 1994) also grow on DMS as sole carbon and energy sources. Roseobacter strains break down DMS into MeSH (Gonza´lez et al., 1999), although in only one case has it been demonstrated that DMS sustains growth (Schaefer et al., 2002). Some of these bacterial isolates are obligate methylotrophs and others can use compounds other than C1; in other cases, they are both methylotrophs and inorganic sulfur oxidizers. Nevertheless, the role that the transformation of DMS by bacterioplankton plays in the ocean is not known. Enrichment with DMS selectively stimulated the growth of Methylophaga (Scha¨fer, 2007; Vila-Costa et al., 2006). Vila-Costa et al. (2006) enriched for Methylophaga in sea water samples to which a small amount of DMS had been added and followed its degradation pathway in the presence and absence of additional carbon sources. Even though the communities did not change significantly between the different treatments, the transformation followed a different pathway when an additional readily utilizable carbon source was present in the medium. S-DMS was completely mineralized to sulfate in DMS enrichments, whilst it was oxidized to DMSO and was not further degraded when glucose was present in the medium. Methylophaga is known to be an obligate methylotroph, such is the case of Methylophaga sulfidovorans, which grows on DMS (deZwart et al., 1996). Although to a lesser extent Thiomicrospira, a sulfur oxidizer, also dominated the communities in these enrichments. Neither Methylophaga nor Thiomicrospora are known to dominate the bacterial communities

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of the ocean and only five Thiomicrospira 16S rRNA sequences were found in the GOS metagenome (Rusch et al., 2007), whereas Methylophaga-related sequences were not detected. Therefore, it is likely that additional, currently unrecognized, groups of bacteria are involved in the degradation of DMS in the natural environment. The role of facultative autotrophic sulfur oxidizers in the degradation of DMS remains also unknown.

6

Genes for the Transformation of DMS

Hyphomicrobium and Thiobacillus both possess an NADH dependent DMS monooxygenase, whose products are MeSH and formaldehyde respectively (de Bont et al., 1981; Kanagawa and Kelly, 1986; Suylen and Kuenen, 1986). A MeSH oxidase participates in the breakdown of MeSH into formaldehyde, sulfide and H2O2 as first reported in Hyphomicrobium (Suylen et al., 1987). No gene that encodes these enzymes has been fully characterized. However, the large subunit of a methanol monooxygenase was induced during growth on DMS by a Methylophaga strain that can grow on DMS as sole carbon source, suggesting that it has DMS monooxygenase activity (Scha¨fer, 2007). Homologs of the gene for this enzyme are found in seven Roseobacter genomes, including that of Sagittula stellata, which is known to metabolize DMS (Gonza´lez et al., 1999), as well as in several strains of Rhodobacter sphaeroides. None of the available SAR11 genomes contain homologs of this putative DMS monooxygenase gene. An alternative to the degradation of DMS exists in which MeSH is not a degradation product (> Fig. 3). DMS is instead oxidized first to DMSO and then further to DMSO2. DMSO2 is broken down into formaldehyde and MSA in Acinetobacter sp. 20B (Horinouchi et al., 1997) and Pseudomonas putida DS1 (Endoh et al., 2003). Genes involved in the oxidation of DMS to DMSO2 have been identified in Acinetobacter sp. 20B. The gene cluster had homology with multicomponent phenol and methane monooxygenases (Horinouchi et al., 1997). A gene cluster for the oxidation of MSA has been described in the soil bacterium Methylosulfonomonas methylovora (de Marco et al., 1999) and the Roseobacter strain Marinosulfonomonas methylotropha (Baxter et al., 2002). The cluster msmABCD codifies an NADHdependent MSA monooxygenase in these two organisms as well as other MSA utilizers.

7

Bacteria Make the Most of DMS(P): Oxidation of Inorganic Sulfur

The reduction state of the atom of sulfur in DMSP or DMS should allow for potentially higher growth yields if an organism is also able to oxidize it. This is supported by the fact that bacteria that both degrade DMS and oxidize inorganic sulfur have been isolated with relative ease (deZwart et al., 1996; Scha¨fer, 2007; Suylen et al., 1986). The genetics of inorganic sulfur oxidation to sulfate has been studied most extensively in lithoautotrophic bacteria (Friedrich et al., 2001), although little is known about the mechanism of oxidation from sulfur to sulfite. Both lithoautotrophs and phototrophs, as well as heterotrophic sulfur oxidizers, share components. Autotrophic sulfur oxidizers of the genus Thiomicrospira were also enriched in DMS-treated mesocosms (Scha¨fer, 2007; Vila-Costa et al., 2006). These organisms could gain additional energy from the inorganic sulfur moiety of DMS during growth on DMS. Reductants and ATP from the oxidation of inorganic sulfur

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can be used to fix formaldehyde released from the degradation of this type of methylated sulfur compounds. Although sulfur oxidizer-related 16S rRNA genes, such as from Thiomicrospira, are not typically found in environmental sequences from the oceanic environment (except for symbiotic communities), the soxB gene, one of the components of the sulfur oxidation system, was found in proportions that reached as high as 25% of the bacterioplankton from the Sargasso Sea (Moran et al., 2004). DMS(P) transformations might account for the widespread occurrence of sulfur oxidation genes in marine bacteria. Inorganic sulfur oxidation has not been as deeply studied in heterotrophic bacteria. sox gene systems appear widespread in members of the marine Roseobacter group (Moran et al., 2007). Analysis of twenty Roseobacter genome sequences reveals 13 that contain the sox operon for sulfur oxidation. Interestingly, isolates of this clade were the first marine strains found to simultaneously possess two key pathways for the degradation of DMSP (Gonza´lez et al., 1999), suggesting this group of marine bacteria are adept at a variety of sulfur transformations.

8

Research Needs

The biogeochemical significance of DMS(P) transformations has been recognized since the 1970s. However, our understanding of the function and importance of DMSP and related compounds in marine food webs has improved significantly in just the last few years. What seems clear is that DMSP and DMS are remarkably versatile substrates when it comes to biological transformations in seawater. Descriptions of key genes and the enzymology of proteins involved in the assimilation and degradation of DMSP have only just begun; however, the little genetics that is known and the distribution of the few genes that have been found in metagenomic libraries are changing our perspective of the roles of DMS(P) in the ocean. Continued study of DMSP and DMS transformations will undoubtedly reveal some of the strategies employed by marine bacteria to adapt to their natural environment and how these adaptations affect cycling of Earth’s major elements.

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De Bont JAM, van Dijken JP, Harder W (1981) Dimethyl sulphoxide and dimethyl sulphide as a carbon, sulphur and energy source for growth of Hyphomicrobium S. J Gen Microbiol 127: 315–323. De Marco P, Moradas-Ferreira P, Higgins TP, McDonald I, Kenna EM, Murrell JC (1999) Molecular analysis of a novel methanesulfonic acid monooxygenase from the methylotroph Methylosulfonomonas methylovora. J Bacteriol 181: 2244–2251. De Souza MP, Yoch DC (1996) N-terminal amino acid sequences and comparison of DMSP lyases from Pseudomonas doudoroffii and Alcaligenes strain M3A. In: Environmental and biological chemistry on dimethylsulfoniopropionate and related sulfonium compounds, RP Kiene, PT Visscher, MD Keller, GO Kirst (eds.). New York: Plenum Press, pp. 293–304. Del Valle DA, Kieber DJ, Kiene RP (2007) Depthdependent fate of biologically-consumed dimethylsulfide in the Sargasso Sea. Mar Chem 103: 197–208. DeZwart JMM, Nelisse PN, Kuenen JG (1996) Isolation and characterization of Methylophaga sulfidovorans sp. nov.: an obligately methylotrophic, aerobic, dimethylsulfide oxidizing bacterium from a microbial mat. FEMS Microbiol Ecol 20: 261–270. Endoh T, Kasuga K, Horinouchi M, Yoshida T, Habe H, Nojiri H, Omori T (2003) Characterization and identification of genes essential for dimethyl sulfide utilization in Pseudomonas putida strain DS1. Appl Microbiol Biotechnol 62: 83–91. Friedrich CG, Rother D, Bardischewsky F, Quentmeier A, Fischer J (2001) Oxidation of reduced inorganic sulfur compounds by bacteria: emergence of a common mechanism? Appl Environ Microbiol 67: 2873–2882. Fuse H, Takimura O, Murakami K, Yamaoka Y, Omori T (2000) Utilization of dimethyl sulfide as a sulfur source with the aid of light by Marinobacterium sp. strain DMS-S1. Appl Environ Microbiol 66: 5527–5532. Gonza´lez JM, Kiene RP, Moran MA (1999) Transformation of sulfur compounds by an abundant lineage of marine bacteria in the a-subclass of the class Proteobacteria. Appl Environ Microbiol 65: 3810–3819. Hanlon SP, Holt RA, Moore GR, McEwan AG (1994) Isolation and characterization of a strain of Rhodobacter sulfidophilus: a bacterium which grows autotrophically with dimethylsulfide as electron-donor. Microbiology 140: 1953–1958. Horinouchi M, Kasuga K, Nojiri H, Yamane H, Omori T (1997) Cloning and characterization of genes encoding an enzyme which oxidizes dimethyl sulfide in Acinetobacter sp. strain 20B. FEMS Microbiol Lett 155: 99–105.

Howard EC, Henriksen JR, Buchan A, Reisch CR, Bu¨rgmann H, Welsh R, Ye W, Gonza´lez JM, Mace K, Joye SB, Kiene RP, Whitman WB, Moran MA (2006) Bacterial taxa that limit sulfur flux from the ocean. Science 314: 649–652. Howard EC, Sun S, Biers EJ, Moran MA (2008) Abundant and diverse bacteria involved in DMSP degradation in marine surface waters. Environ Microbiol 10: 2397–2410. Kanagawa T, Kelly DP (1986) Breakdown of dimethyl sulfide by mixed cultures and by Thiobacillus thioparus. FEMS Microbiol Lett 34: 13–19. Kettle AJ, Andreae MO (2000) Flux of dimethylsulfide from the oceans: a comparison of updated data sets and flux models. J Geophys Res 26: 26793–26808. Kiene RP, Hoffman Williams LP, Walker JE (1998) Seawater microorganisms have a high affinity glycine betaine uptake system which also recognizes dimethylsulfoniopropionate. Aquat Microb Ecol 15: 39–51. Kiene RP, Linn LJ (2000) Distribution and turnover of dissolved DMSP and its relationships with bacterial production and dimethylsulfide in the Gulf of Mexico. Limnol Oceanogr 45: 849–861. Kiene RP, Linn LJ, Bruton JA (2000) New and important roles for DMSP in marine microbial communities. J Sea Res 43: 209–224. Kiene RP, Linn LJ, Gonza´lez JM, Moran MA, Bruton JA (1999) Dimethylsulfoniopropionate and methanethiol are important precursors of methionine and protein-sulfur in marine bacterioplankton. Appl Environ Microbiol 65: 4549–4558. Kiene RP, Oremland RS, Catena A, Miller LG, Capone DG (1986) Metabolism of reduced methylated sulfur compounds in anaerobic sediments and by a pure culture of an estuarine methanogen. Appl Environ Microbiol 52: 1037–1045. Malmstrom RR, Kiene RP, Cottrell MT, Kirchman DL (2004) Contribution of SAR11 bacteria to dissolved dimethylsulfoniopropionate and amino acid uptake in the North Atlantic ocean. Appl Environ Microbiol 70: 4129–4135. Moran MA, Belas R, Schell MA, Gonza´lez JM, Sun F, Sun S, et al. (2007) Ecological genomics of marine roseobacters. Appl Environ Microbiol 73: 4559–4569. Moran MA, Buchan A, Gonza´lez JM, Heidelberg JF, Whitman WB, Kiene RP, et al. (2004) Genome sequence of Silicibacter pomeroyi reveals adaptations to the marine environment. Nature 432: 910–913. Omori T, Saiki Y, Kasuga K, Kodama T (1995) Desulfurization of alkyl and aromatic sulfides and sulfonates by dibenzothiophene-desulfurizing Rhodococcus sp. strain SY1. Biosci Biotechnol Biochem 59: 1195–1198.

Genetics and Molecular Features of Bacterial DMSP and DMS Transformations Pol A, Op den Camp HJ, Mees SG, Kersten MA, van der Drift C (1994) Isolation of a dimethylsulfideutilizing Hyphomicrobium species and its application in biofiltration of polluted air. Biodegradation 5: 105–112. Rusch DB, Halpern AL, Sutton G, Heidelberg KB, Williamson S, Yooseph S, et al. (2007) Oceanic metagenomics: the Sorcerer II global ocean sampling expedition: northwest Atlantic through eastern tropical Pacific. PLoS Biol 5: e77. Schaefer JF, Goodwin KD, McDonald IR, Murrell JC, Oremland RS (2002) Leisingera methylohalidivorans gen. nov., sp. nov., a marine methylotroph that grows on methyl bromide. Int J Syst Evol Microbiol 52: 851–859. Scha¨fer H (2007) Isolation of Methylophaga spp. from marine dimethylsulfide-degrading enrichment cultures and identification of polypeptides induced during growth on dimethylsulfide. Appl Environ Microbiol 73: 2580–2591. Simo´ R (2001) Production of atmospheric sulfur by oceanic plankton: biogeochemical, ecological and evolutionary links. Trends Ecol Evol 16: 287–294. Simo´ R, Archer SD, Pedro´s-Alio´ C, Gilpin L, StelfoxWiddicombe CE (2002) Coupled dynamics of dimethylsulfoniopropionate and dimethylsulfide cycling and the microbial food web in surface waters of the North Atlantic. Limnol Oceanogr 47: 53–61. Simo´ R, Pedro´s-Alio´ C (1999) Short-term variability in the open ocean cycle of dimethylsulfide. Global Biogeochem Cycles 13: 1173–1181. Suylen GMH, Kuenen JG (1986) Chemostat enrichment and isolation of Hyphomicrobium EG a dimethyl sulfide oxidizing methylotroph and reevaluation of Thiobacillus MS1. Antonie Leeuwenhoek 52: 281–293. Suylen GMH, Large PJ, van Dijken JP, Kuenen JG (1987) Methyl mercaptan oxidase, a key enzyme in the metabolism of methylated sulphur compounds by Hyphomicrobium EG. J Gen Microbiol 133: 2989–2997. Suylen GMH, Stefess GC, Kuenen JG (1986) Chemolithotrophic potential of a Hyphomicrobium species, capable of growth on methylated sulphur compounds. Arch Microbiol 146: 192–198.

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Tanimoto Y, Bak F (1994) Anaerobic degradation of methylmercaptan and dimethyl sulfide by newly isolated thermophilic sulfate-reducing bacteria. Appl Environ Microbiol 60: 2450–2455. Thompson AS, Owens NJP, Murrell JC (1995) Isolation and characterization of methanesulfonic aciddegrading bacteria from the marine environment. Appl Environ Microbiol 61: 2388–2393. Todd JD, Rogers R, Li YG, Wexler M, Bond PL, Sun L, Curson AR, Malin G, Steinke M, Johnston AW (2007) Structural and regulatory genes required to make the gas dimethyl sulfide in bacteria. Science 315: 666–669. Tripp JH, Kitner JB, Schwalbach MS, Dacey JWH, Wilhelm LJ, Giovannoni SJ (2008) SAR11 marine bacteria require exogenous reduced sulphur for growth. Nature 452: 741–744. Vallino JJ, Hopkinson CS, Hobbie JE (1996) Modeling bacterial utilization of dissolved organic matter: optimization replaces Monod growth kinetics. Limnol Oceanogr 41: 1591–1609. Vila M, Simo´ R, Kiene RP, Pinhassi J, Gonza´lez JM, Moran MA, Pedro´s-Alio´ C (2004) Use of microautoradiography combined with fluorescence in situ hybridization to determine dimethylsulfoniopropionate incorporation by marine bacterioplankton taxa. Appl Environ Microbiol 70: 4648–4657. Vila-Costa M, del Valle DA, Gonza´lez JM, Slezak D, Kiene RP, Sa´nchez O, Simo´ R (2006) Phylogenetic identification and metabolism of marine dimethylsulfideconsuming bacteria. Environ Microbiol 8: 2189–2200. Vila-Costa M, Pinhassi J, Alonso C, Pernthaler J, Simo´ R (2007) An annual cycle of dimethylsulfoniopropionate-sulfur and leucine assimilating bacterioplankton in the coastal NW Mediterranean. Environ Microbiol 9: 2451–2463. Visscher PT, Taylor BF (1994) Demethylation of dimethylsulfoniopropionate to 3-mercaptopropionate by an aerobic marine bacterium. Appl Environ Microbiol 60: 4617–4619. Yoch DC (2002) Dimethylsulfoniopropionate: its sources, role in the marine food web, and biological degradation to dimethylsulfide. Appl Environ Microbiol 68: 5804–5815.

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28 Environmental Mining of Biological Activities on Hydrocarbons K. Watanabe Research Center for Advanced Science and Technology, University of Tokyo, Tokyo, Japan [email protected] 1 PCR-Based Approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1214 2 Metagenome-Library Approaches . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1216 3 Conclusions and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1217

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Abstract: Microbes that utilize hydrocarbons for their growth were recognized more than 100 years ago. ZoBell (1946) reviewed knowledge of the time concerning the action of microorganisms on hydrocarbons; according to his review, Miyoshi found in 1895 that thin layers of paraffin were penetrated by Botrytis cinerea. In addition, it was also introduced in that review article that the ability of fungi to attack paraffin had been reported in 1906, showing that various soil molds could decompose paraffin and utilize it as a sole source of energy. Since then, numerous studies have been performed to isolate and cultivate microbes that are able to degrade and transform hydrocarbons (Atlas, 1981). Although a large part of these studies have focused on aerobic microbes that utilize molecular oxygen as the respiratory electron acceptor and as agents to oxidize hydrocarbons, recent studies have also shown that microbes can degrade hydrocarbons under various anaerobic conditions. These microbes include nitratereducing bacteria, sulphate-reducing bacteria, and iron-reducing bacteria (Heider et al., 1998). In addition, it has been also reported that long-chain alkanes were decomposed to methane under anaerobic fermentative conditions (Zengler et al., 1999). Since these microbes do not utilize molecular oxygen to oxidize hydrocarbons, their molecular mechanisms to transform hydrocarbons are entirely different from those in aerobic counterparts. These activities of microbes are of industrial interests in terms of their application to bioremediation and chemical synthesis. Hydrocarbons are the major components of petroleum and fuels that widely contaminate the environment as a result of human activities. Bioremediation that utilize in situ or artificially introduced hydrocarbon-degrading microbes is considered to be an environmentally friendly and cost-saving option to clean up contaminated sites (Leahy and Colwellm, 1990). In addition, hydrocarbon-transforming enzymes (e.g., oxygenases) in these microbes are being exploited in various chemical processes (Schmid et al., 2001), since they allow the highly chemo-, regio-, and enantioselective functionalization of hydrocarbons under mild conditions (Bu¨hler and Schmid, 2004). It is essential for these efforts to obtain microbes and enzymes that efficiently and/or specifically transform certain species of hydrocarbons. The natural environment is the source of useful microbes and enzymes. As described above, microbes that have hydrocarbon-degrading activities have been isolated, and genes coding for degradative enzymes have subsequently been cloned from these microbes. This scheme is conventionally employed to fish out desired biological activities from the natural environment, although we now realize that only a small fraction of natural microbes can be cultivated and isolated by the conventional microbiological technique (Amann et al., 1995). In order to access to more diverse enzymes present in uncultured microbes, scientists have started to use molecular and genomic techniques for screening of genome mixtures directly extracted from natural microbial communities (i.e., metagenomes) (Handelsman et al., 1998). In this chapter, I summarize recent progresses on environmental mining of biological activities on hydrocarbons. In particular, we describe successful applications of PCR-based and metagenomic approaches for obtaining novel genes coding for hydrocarbon-transforming enzymes.

1

PCR-Based Approaches

A large number of microbes capable of degrading different types of pollutants have been isolated, and genes coding for various hydrocarbon-degrading enzymes have been cloned and sequenced. Based on sequence information obtained in these studies, it is possible to design PCR primers that can be used for detecting degradative genes from microorganisms in the

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environment. PCR primers that have been successfully used for such purposes have recently been summarized elsewhere (Watanabe and Kasai, 2007). These primers are designed by comparing nucleotide sequences of genes or amino-acid sequences of enzymes in a same functional group; namely, alignment of sequences allows identification of conserved regions whose sequences can be used to design PCR primers. These PCR primers are potentially useful to detect and clone novel genes for hydrocarbon transformation, although we should be aware of the fact that they can amplify partial fragments (central parts) that are in most cases insufficient to synthesize functional enzymes. Several approaches are possible to construct an entire gene. First, Okuta et al. (1998) demonstrated that environmental PCR products can be connected to 3´ and 5´ sequences of a known gene to construct a functional hybrid gene. They isolated central segments of catechol 2, 3-dioxygenase (C23O) genes from environmental samples by PCR using primers that were designed from conserved sequences among a wide range of C23O genes and were connected to restriction sites, followed by PCR products being inserted between 5´ and 3´ regions of nahH (the structural gene for C23O encoded by catabolic plasmid NAH7). More than 90% of the hybrid C23O genes expressed the activity, showing that the method is useful to create, without isolating bacteria, a library of functional hybrid genes. In order to obtain authentic franking regions of an environmental PCR product, a method (IAN-PCR) has been developed by modifying the conventional inverse PCR technique (Uchiyama and Watanabe, 2006). When a PCR product is derived from a low copy-number metagenome fragment, franking regions cannot generally be obtained by the standard inverse PCR scheme due to low amplification efficiency. IAN-PCR was designed to improve the amplification efficiency and included the following steps (> Fig. 1): (1) inverse PCR in which one primer is connected to an affinity tag; (2) affinity purification of PCR products for removing background metagenome; and (3) nested PCR to recover target flanking regions (IAN-PCR). In a model experiment of IAN-PCR, flanking regions of a gene fragment in

. Figure 1 Inverse nested PCR (IN-PCR) and inverse affinity nested PCR (IAN-PCR) for amplifying franking regions of target genes from environmental metagenomes.

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Ralstonia eutropha were recovered from mixtures of Ralstonia and Escherichia genomes by standard inverse PCR, inverse PCR coupled to nested PCR (IN-PCR), and IAN-PCR, showing that they were recovered when ratios of Ralstonia genome to the background Escherichia genome were greater than 10 1, 10 3, and 10 5, respectively. Thus IAN-PCR was considered to applicable to recover franking regions from 10,000-time less metagenome fragments. Another approach to apply inverse PCR to low copy-number metagenome fragments has been reported recently, in which rolling-circle amplification was used with a site-specific primer containing locked nucleic acids (Yamada et al., 2008).

2

Metagenome-Library Approaches

A metagenome-library approach includes following steps: (1) isolation of mixed genomic DNA from an environmental sample, (2) cloning of digested metagenome fragments into a suitable vector, (3) transformation of a host bacterium, and (4) analysis or screening of transformant clones. The clones obtained can be subjected to shotgun sequencing and/or functional screening for acquired traits, such as enzyme activity and/or antibiotic production. One example of the functional screening is presented by Van Hellemond et al. (2007), in which they attempted to obtain a type of oxygenases that are able to form the blue pigment indigo when it is expressed in Escherichia coli. Using this approach, they identified a novel oxygenase from a metagenomic library derived from loam soil. This oxygenase shows 50% sequence identity to styrene monooxygenases from pseudomonads (StyA). The newly identified monooxygenase catalyzes the epoxidation of styrene and styrene derivatives and forms the corresponding (S)-epoxides with excellent enantiomeric excess and therefore is named styrene monooxgenase subunit A (SmoA). SmoA shows high enantioselectivity towards aromatic sulfides. This excellent enantioselectivity in combination with the moderate sequence identity forms a clear indication that SmoA from a metagenomic origin represents a new enzyme within the small family of styrene monooxygenases. In another example, a metagenomic library was constructed from activated sludge used to treat coke plant wastewater (Suenaga et al., 2007). Metagenome fragments were cloned into fosmids, and the resulting Escherichia coli library was screened for extradiol dioxygenases (EDOs) using catechol as a substrate (yellow intermediate metabolite [2-hydroxymuconate semialdehyde] produced from catechol is checked to identify the catabolic activity), yielding 91 EDO-positive clones. Based on their substrate specificity for various catecholic compounds, 38 clones were subjected to sequence analysis. Each insert contained at least one EDO gene, and a total of 43 EDO genes were identified. More than half of these belonged to new EDO subfamilies: I.1.C (2 clones), I.2.G (20 clones), I.3.M (2 clones) and I.3.N (1 clone). The fact that novel I.2.G family genes were over-represented in these clones suggested that these genes play a specific role in environmental aromatic degradation. The I.2.G clones were further classified into six groups based on single-nucleotide polymorphisms (SNPs). An advantage of these functional screening is that a metagenome fragment containing an entire gene fragment can be obtained. However, we should know that efficient screening is possible (as exemplified by the above cases) only if easily detectable compounds (such as colored compounds) are produced. In addition, a recent study has developed a gene-expression dependent screening of a metagenome library, i.e., substrate induced gene expression screening (SIGEX), for isolating novel catabolic genes (Uchiyama et al., 2005; Uchiyama and Watanabe, 2008). This method is entirely different from the above-mentioned screening methods, since it is not dependent on nucleotide sequences and enzyme functions. In this method, an operon-trap gfp-expression

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vector is used for constructing a metagenomic library, which enables subsequent high throughput screening for GFP-expressed positive clones with the aid of fluorescence-activated cell sorting. This was applied to fishing out operon fragments involved in aromatic hydrocarbon degradation from a metagenome library constructed from oil-contaminated groundwater (Uchiyama et al., 2005). It has been shown that, in metagenome fragments isolated by SIGEX with benzoate as an induction substrate, open reading frames homologous to know catabolic genes were highly enriched, and some of them were substantially similar (over 70%) to genes in the known benzoate-degradative pathway. A novel cytochrome P450 converting 4-hydroxybenzoate to protocatechuate was identified; an amino-acid sequence of this enzyme was very different from known P450 sequences, demonstrating the power of the SIGEX scheme. Merits and demerits of the SIGEX scheme in comparison with other screening methods has been discussed previously (Uchiyama and Watanabe, 2007). One of common problems associated with the metagenome approach is that we are difficult to access genomes of minor species, even though they include a vast majority of the natural genetic diversity. In order to obviate this limitation, Podar et al. (2007) used phylogenetically directed cell separation by fluorescence in situ hybridization and flow cytometry, followed by amplification and sequencing of a fraction of the genomic DNA of several bacterial cells that constituted a minor population in a soil sample. They succeeded in partial reconstruction of the minority bacterium (the TM7 bacterium), suggesting that this approach is useful to access any specific members of a community and an alternative way to assess the community’s metabolic potential. Similarly, if one removes cells of several major populations from an environmental sample by that scheme, a possibility to access minor populations can be increased.

3

Conclusions and Research Needs

This chapter introduced molecular and metagenomic methods that are applicable to fish out genes for novel hydrocarbon-transforming enzymes from environmental samples. It is important to realize that each of these methods has its own territory (in terms of the types of genes they can fish out); namely, none of them is able to access to a broad range of genes in the environment. For example, PCR-mediated methods are accessible to genes in the environment that are similar (in nucleotide sequences) to genes whose sequences were used to design PCR primers. We therefore need to understand the capacity of each method and select an appropriate method that is suitable for a particular purpose. In addition, the development of a new screening strategy should be the key to accessing hitherto unexploited genes with a high degree of novelty.

References Amann RI, Ludwig W, Schleifer KH (1995) Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol Rev 59: 143–169. Atlas RM (1981) Microbial degradation of petroleum hydrocarbons: an environmental perspective. Microbiol Rev 45: 180–209. Bu¨hler B, Schmid A (2004) Process implementation aspects for biocatalytic hydrocarbon oxyfunctionalization. J Biotechnol 113: 183–210.

Handelsman J, Rondon MR, Brady SF, Clardy J, Goodman RM (1998) Molecular biological access to the chemistry of unknown soil microbes: a new frontier for natural products. Chem Biol 5: R245–R249. Heider J, Spormann AM, Beller HR, Widdel F (1998) Anaerobic bacterial metabolism of hydrocarbons. FEMS Microbiol Rev 22: 459–473. Leahy JG, Colwell RR (1990) Microbial degradation of hydrocarbons in the environment. Microbiol Rev 54: 305–315.

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Okuta A, Ohnishi K, Harayama S (1998) PCR isolation of catechol 2,3-dioxygenase gene fragments from environmental samples and their assembly into functional genes. Gene 212: 221–228. Podar M, Abulencia CB, Walcher M, Hutchison D, Zengler K, Garcia JA, Holland T, Cotton D, Hauser L, Keller M (2007) Targeted access to the genomes of low-abundance organisms in complex microbial communities. Appl Environ Microbiol 73: 3205–3214. Schmid A, Dordick JS, Hauer B, Kiener A, Wubbolts M, Witholt B (2001) Industrial biocatalysis today and tomorrow. Nature 409: 258–268. Suenaga H, Ohnuki T, Miyazaki K (2007) Functional screening of a metagenomic library for genes involved in microbial degradation of aromatic compounds. Environ Microbiol 9: 2289–2297. Uchiyama T, Abe T, Ikemura T, Watanabe K (2005) Substrate-induced gene-expression screening of environmental metagenome libraries for isolation of catabolic genes. Nat Biotechnol 23: 88–93. Uchiyama T, Watanabe K (2006) Improved inverse PCR scheme for metagenome walking. Biotechniques 41: 183–188. Uchiyama T, Watanabe K (2007) The SIGEX scheme: high throughput screening of environmental

metagenomes for the isolation of novel catabolic genes. Biotechnol Genet Eng Rev 24: 107–116. Uchiyama T, Watanabe K (2008) Substrate-induced gene-expression (SIGEX) screening of metagenome libraries. Nat Protocol 3: 1202–1212. Van Hellemond EW, Janssen DB, Fraaije MW (2007) Discovery of a novel styrene monooxygenase originating from the metagenome. Appl Environ Microbiol 73: 5832–5839. Watanabe K, JKasai Y (2007) Emerging technologies to analyse natural attenuation and bioaugmentation. In Microbial Bioremediation: Genomics and Molecular Biology. Cambridge, UK: Caister Academic Press, pp. 295–316. Yamada K, Terahara T, Kurata S, Yokomaku T, Tsuneda S, Harayama S (2008) Retrieval of entire genes from environmental DNA by inverse PCR with preamplification of target genes using primers containing locked nucleic acids. Environ Microbiol 10: 978–987. Zengler K, Richnow HH, Rossello´-Mora R, Michaelis W, Widdel F (1999) Methane formation from longchain alkanes by anaerobic microorganisms. Nature 401: 266–269. ZoBell CE (1946) Action of microorganisms on hydrocarbons. Bacteriol Rev 10: 1–49.

29 Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements R. R. Fulthorpe1 . E. M. Top2 Physical and Environmental Sciences, University of Toronto at Scarborough, Toronto, ON, Canada [email protected] 2 Department of Biological Sciences, Initiative for Bioinformatics and Evolutionary Studies (IBEST), University of Idaho, Moscow, ID, USA [email protected]

1

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1220

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Catabolic Mobile Elements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1220

3 3.1 3.1.1 3.1.2 3.1.3 3.1.4 3.1.5 3.1.6 3.1.7

Examples of Degradation Pathways Most Likely Evolved Through Assembly by Means of Plasmids and Transposons . . . . . . . . . . . . . . . . . . . 1221 Aromatic Compounds, a Brief Summary . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1222 Pentachlorophenol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1222 Nitrobenzene . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1222 Atrazine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1226 TSA – Toluenesulfonate . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1227 Nylon Oligomers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1228 2,4-Dichlorophenoxacetic Acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1228 Carbazole . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1229

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1229

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Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

Abstract: There is no doubt today that the evolution of complex catabolic pathways for the degradation of xenobiotic compounds in bacteria is mediated by mobile genetic elements, such as transposons, insertion sequences, integrative and conjugative elements (ICE), and plasmids. Genetic analysis of several such pathways strongly suggests that the genes that encode the various enzymatic steps are derived from existing single genes or whole operons, which are brought together by inter- and intra-cellular gene transfer mechanisms. Subsequently, selection acts on specific mutations that result in more optimal enzymatic activity for specific recalcitrant compounds. This clever system of ‘‘patchwork assembly’’ allows bacterial communities to rapidly adapt to new xenobiotics. Those bacteria that have assembled successful degradation pathways through this approach are the ones we find today in various polluted habitats. This chapter gives a brief overview of several examples of successful in situ pathway constructions that have been selected over evolutionary time.

1

Introduction

Bacteria regularly encounter xenobiotics in their natural habitats, i.e., compounds that have been released into the environment by the action of humans (Leisinger, 1983). While acquiring the ability to use such novel compounds as carbon or nitrogen sources may seem a long and difficult process, it is often not an insurmountable task for the bacteria given that these molecules are often structurally related to naturally occurring compounds, such as metabolites produced by plants and microorganisms (Myneni, 2002). Thus rather than evolving enzymes ‘‘from scratch,’’ a small number of evolutionary steps can sometimes be sufficient for a bacterial population to acquire the ability to degrade a novel compound. There are basically two kinds of such evolutionary steps: (i) genetic changes within the genome, such as point mutations or short insertions or deletions, and transposition and recombination events, which can result in rearrangements, changes in gene regulation, deletions or duplications; and (ii) acquisition of genetic information from related or phylogenetically distinct populations by horizontal or lateral gene transfer (HGT, LGT) through conjugation by means of plasmids or other conjugative elements, transformation, or transduction (Liu and Sulfita, 1993). While HGT allows for the wholesale uptake of genes, a form of ‘‘evolution in quantum leaps’’ (Hacker and Carniel, 2001), optimal biodegradation of a new compound may not be achieved by simply recruiting different genes, but subsequent genetic changes may be required. Eventually, the individuals best able to degrade the xenobiotic and use it as a source of carbon or nitrogen under the extant environmental conditions, will outcompete others and constitute the bacterial strains and biochemical pathways that we know about today. We can think of this ‘‘patchwork assembly’’ as natural tinkering with genetic material, where the catabolic pathway that provides the largest fitness benefit to the bacterial host is selected over evolutionary time.

2

Catabolic Mobile Elements

Before we can understand pathway assembly we need to know the main agents of horizontal gene transfer that bring about the mosaic pathways we find today, the mobile genetic elements (MGE). Among the transposons that carry determinants for organic pollutant degradation there are two large groups: class I and class II transposons, often found on plasmids (Tsuda et al., 1999). Of the insertion sequences that flank catabolic genes, IS1071-like sequences are a

Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

29

notorious example, found adjacent to a long list of catabolic genes in a diverse range of bacteria (Di Gioia et al., 1998; Rousseaux et al., 2002). The frequent presence of this IS near catabolic genes and its absence on most drug resistance plasmids (except for the IncP-1 multidrug resistance plasmid pB10, (Schlu¨ter et al., 2003)) is puzzling and fodder for future studies. In addition to these MGE there is a growing range of catabolic genetic elements that have been shown to be mobile or are potentially mobile, such as clc, bph-sal, and Tn4371, the so-called integrative and conjugative elements (ICE) (Burrus et al., 2002; Gaillard et al., 2006; Ravatn et al., 1998; Toussaint et al., 2003; Van Der Meer and Sentchilo, 2003). One of the major modes of transport of catabolic genes between strains of related or phylogenetically distinct species is conjugation by means of plasmids. Catabolic plasmids, that encode degradation of man-made or natural organic compounds, are usually large (>50 kb), because several enzymes are required for degradation and because most plasmids are self-transferable. So far, catabolic genes in Gram negative bacteria have been found only on six groups of plasmids, i.e., IncP-1, IncP-2, IncP-7, and IncP-9 plasmids mostly found in Pseudomonas and b-Proteobacteria, unclassified plasmids from the a-Proteobacteria (mainly sphingomonads) and Flavobacterium (Nojiri et al., 2004; Ogawa et al., 2004; Sota and Top, 2008; Top et al., 2000; Williams et al., 2004). In Gram positive bacteria, catabolic plasmids are mainly found in Arthrobacter and Rhodococcus (Ogawa et al., 2004). It is intriguing that the plasmids that encode degradation of naturally occurring hydrocarbons such as naphthalene and toluene almost always belong to the IncP-2, IncP-7, and IncP-9 groups (incompatibility groups defined in Pseudomonas), while those involved in the degradation of man-made compounds such as several chlorinated organic compounds are typically IncP-1 (=IncP in E. coli plasmid classification) and a-proteobacterial plasmids (Mergeay et al., 1990; Nojiri et al., 2004; Ogawa et al., 2004; Top et al., 2000). We have previously speculated that there may be a correlation between the high promiscuity and transfer rates of IncP-1 plasmids and their ability to acquire genes encoding degradation of recently introduced xenobiotics. For more details about some of the catabolic plasmids and their role in pathway evolution, we refer to van der Meer et al. (2007) and Williams et al. (2004).

3

Examples of Degradation Pathways Most Likely Evolved Through Assembly by Means of Plasmids and Transposons

The currently accepted hypothesis is that the genes used in the assembly of pathways for the degradation of novel, xenobiotic compounds are derived from existing genes or whole operons that are brought together by the MGE described above. This assembly of genes from different sources has been referred to as a ‘‘mosaicity’’ (Maeda et al., 2003) or a ‘‘patchwork’’ (Copley, 2000). The evidence in support of this hypothesis lies in the many examples of degradative organisms that are carrying genes whose products culminate in successful degradation of novel compounds. In the initial stages of pathway development, the genes can be found in different species or strains, on different elements in the same strains (plasmids or genomes), widely spaced on the same element, under separate regulation, and frequently associated with IS elements or transposons. Where the genetic locations of genes are not known (i.e. for PCP degraders), the disparate sources of the component genes or operons are evident in their sequence similarity to genes from different organisms, or in the differing GC contents of the pathway genes. To illustrate this, we describe just some of the pathways found in bacteria that degrade hydrocarbons or their derivatives.

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Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

Aromatic Compounds, a Brief Summary

The classical example of patchwork assembly is the aerobic degradation of aromatic compounds, which typically requires a few steps to channel the parent compounds toward a few dihydroxylated intermediates (van der Meer et al., 1992; Jimenez et al., 2004; Pieper and Reineke, 2004). Degradation of compounds such as benzoate, benzene, phenol, often occurs through catechol as the central intermediate. This aromatic ring is then opened through either intradiol cleavage (ortho-cleavage) by a catechol 1,2-dioxygenase (C12O) or extradiol cleavage (meta-cleavage) by a catechol 2,3-dioxygenase (Jimenez et al., 2004; Pieper and Reineke, 2004). In proteobacteria the enzymes that mediate catechol degradation usually do not allow degradation of chlorosubstituted catechols. Those are often degraded by enzymes of the chlorocatechol pathway, or modified ortho-cleavage pathway, using enzymes that have high activity against the chlorinated intermediates (Reineke, 1998; Reineke and Knackmuss, 1988) (See also > Chapter 5, Vol. 2, Part 2). Thus degradation of a very diverse set of chlorinated aromatic compounds occurs in a very similar fashion. First, in the so-called upper pathway, a few initial enzymatic steps convert the molecule to a central intermediate like chlorocatechol, which is then further degraded through a common modified ortho-cleavage pathway, and steps of the 3-oxoadipate pathway that channel the compounds into the tricarboxylic acid cycle. Many studies have suggested that the genes encoding the first steps were recruited by the bacterial cell from other hosts, mostly mediated by plasmids and transposons, and then underwent mutations that led to more optimal enzymatic reactions (Ca´mara et al., 2007; Top and Springael, 2003; Van Der Meer and Sentchilo, 2003). Below and in > Table 1 we summarize the more recently described examples of pathway assembly.

3.1.1

Pentachlorophenol

Evidence for the involvement of specific mobile genetic elements in the assembly of a pathway for the aerobic degradation of pentachlorophenol is lacking, as no plasmids, IS, transposons or other MGE have been located in these organisms. However an examination of the genes encoding the key degradative steps suggests they are derived from two separate pathways, that for 2,6 dichlorophenol (a natural occurring chloroaromatic compound) and the tyrosine pathway (Copley, 2000). The first gene in the pathway is frequently found alone in diverse organisms (Crawford et al., 2007), supporting the hypothesis that pcpB was recruited from other pathways.

3.1.2

Nitrobenzene

Nitrobenzene degradation proceeds via the partial reduction of the nitro group by a nitrobenzene reduction (nbzA), followed by the action of a mutase gene (nbzB, habB or habA) to produce 2- aminophenol, which undergoes meta ring fission by an aminophenol dioxygenase (nbzCa, nbzCb, or amnB, amnA). These pathways have been studied in three Pseudomonas species, P. pseudoalcaligenes JS45, isolated from soil near a nitrobenzene factory in the US, in P. putida HS12, isolated from contaminated soil in Korea (Johnson and Spain, 2003) and the aminophenol degrading Pseudomonas strain AP-3 (Takenaka, S. et al., 1998; Takenaka, D. et al., 2000). The degradation of nitroaromatics can follow different pathways in different

Study organism

Genes

Toluenesulfonate (Tralau et al., 2001)

Comamonas testosteroni T-2

Pseudomonas sp. ADP Isopropylamino hydrolase N-isopropylammelid, Cyanuric amidohydrolase, Nicotinamidase, Urea amidolyase

atzC atzDEF

psbA(C)

p-sulfobenzoate3,4-dioxygenase

p-sulfobenzaldehyde dehydrogenase

Toluenesulfonate methylmono-oxygenase p-sulfobenzyl alcohol dehydrogenase

Hydroxyatrazine hydrolase

atzB

tsaMBCD

Atrazine chlorohydrolase

N-isopropylammelid isopropylamino hydrolase

pT2L

pTSAa

pADP-1a

IS1071, putative tsaTn, Tn402/ 5090

IS1071, elements of Tn5043, Tn5041, IS801

pAA1a

Hydroxyatrazine hydrolase

nd nd

nd

IS and Tn’s

Triazine hydrolase

Plasmid

Triazine hydrolase

Enzyme

atzA

atzC

Nocardiodes C190 trzN Atrazine (De Souza et al., 1998b; Martinez et al., 2001; Arthrobacter trzN Mulbry et al., 2002; Sajjaphan aurescens TC1 atzB et al., 2004; Topp et al., 2000b)

Compound (References)

. Table 1 A few examples of degradation pathways evolved through gene assembly

Delftia acidovorans Mu4, MoP1, Stenotrophomonas maltophilia MuF, Pseudomonas pseudoalcaligenes, Ralstonia picketti TKR, Unidentified strain TA12, Pseudomonas ISP2

Genes found in Chelatobacter, Stenotrophomonas, Pseudoaminobacter, Agrobacterium, Clavibacter, Rhizobium, Pseudomonas, Alcaligenes, Ralstonia

Same genes in Arthrobacter crystallopoietes

Homologues in

Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

29 1223

Pentachlorophenol (Copley, 2000; Crawford et al., 2007)

Pseudomonas. pseudoalcaligenes JS45

Nitrobenzene (Johnson and Spain, 2003; Park and Kim, 2000; Takenaka et al., 2000)

Sphingobium chlorophenolicum ATCC39723

P. putida HS12

Study organism

Hydroxylamino benzene mutase

habA habB (silent)

Tetrachlorohydroquinone dehalogenase DCHQ dioxygenase

pcpC pcpA

2-aminophenol 1,6 dioxygenase, 2-aminomuconic semialdehyde dehydrogenase

nbzCDE

pcp-4-monooxygenase

Hydroxylamino benzene mutase

nbzB

pcpB

Nitrobenzene reductase

nbzA

nbzCDEFGI

Nitrobenzene reductase

Enzyme

nbzA

Genes

nd

nd

nd

pNB1

pNB2

pNB1

nd

Plasmid

nd

nd

nd

tnpA gene similar to Tn5501

nd

IS and Tn’s

Homologues in

pcpA derived from 2,6 dichlorophenol pathway

pcpC -derived from tyrosine pathway

pcpB – putatively derived from 2,6 dichlorophenol pathway Widespread in degraders/non-degraders

Pseudomonas sp. AP3

Pseudomonas putida ZWL73, Comamonas sp. CNB-1 (pCNB1)

Pseudomonas sp AP3

Pseudomonas putida ZWL73, Comamonas sp. CNB-1 (pCNB1)

29

Compound (References)

. Table 1 (Continued)

1224 Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

Pseudomonas sp. NK87

Sphingomonas KA1

Carbazole (Maeda et al., 2003; Pseudomonas Shintani et al., 2007; Urata sp. CA10 et al., 2006)

Denotes IncP-1b backbone

a

tfdCDEF

Cupriavidus tfdA necator (Ralstonia eutropha) JMP134 tfdB

6-aminohexanote linear dimer hydrolase

nylB pNAD6

pNAD2

pOAD2

pJP4a

Ferrodoxin/reductase

fdrI, fdrII

Similar to above

fdxI

See refs

Anthranilate degradation

Carbazole degradation

carABC

AntABC

pCAR3

carABCDEF Multicomponent enzymes pCAR1

6-aminohexanoate cyclic dimer hydrolase

6 aminohexanoate oligomer hydrolase

6-aminohexanote linear dimer hydrolase

6-aminohexanoate cyclic dimer hydrolase

Orthocleavage chlorocatechol pathway

Phenol monooxygenase

Alpha-keto glutarate dioxygenase

nylA

nylC

Nylon oligomers (Negoro Arthrobacter K172 nylA et al., 1983; Okada et al., 1983; Kanagawa et al., 1989; Kato nylB et al., 1994)

2,4-Dichlorophenoxyacetic acid (Trefault et al., 2004)

Tn4676

IS6100

IS6100

IS1071, ISJP4

Operons found associated with different flanking IS elements in Pseudomonas K23, Janthiniobacterium J3

nylB in Arthobacter aurescens TC1, Bradyrhizobium ORS278

All three genes also found in Agromyces KY5R and Kocuria KY2

tfdCDEF homologues in Pseudomonas sp. strain B13 (clc Island), Burkholderia xenovorans LB400, Bordetella petri DSM12804, Alcaligenes sp st NyZR15, Pseudomonas aeruginosa JB2 among others

tfdA – widespread in degraders, nondegraders

Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

29 1225

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Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

organisms and the notation can be confusing. Comamonas sp. JS765 uses a multicomponent dioxygenase encoded by the genes nbzAa-nbzAd, related to similar genes in Acidovorax sp. JS42 used for dinitrotoluene (Lessner et al., 2002). These must not be confused with the gene nbzA for nitrobenzene reductase. The JS45 genes, encoding the partially reductive pathway, genes are often found in different locations of the genome in the same organism, and have different phylogenetic histories. Strains JS45 and HS12 both carry nbzA genes that are 93% similar. One of these is located on a catabolic plasmid (pNB1) in HS12 (Park and Kim, 2000). However, they use different enzymes for the mutase step. Strain HS12 utilizes NbzB, encoded by a plasmid different from that carrying nbzA (pNB2 rather pNB1), while strain JS45 carries two mutase genes, habA (which it expresses) and habB (which is silent). The genes are only 44% similar, have different GC contents and lie 2.5 kb apart. Curiously habB, silent in strain JS45, is virtually identical to the nbzB gene used by strain HS12. Genes highly similar to nbzA have been found in chloronitrobenzene degraders - in P. putida ZWL73 (Xiao et al., 2006) and Comamonas sp. CNB-1 (pCNB1) (Ma et al., 2007). The action of the nbzA and nbzB or habA gene products culminates in the intermediate aminophenol. The three Pseudomonas strains HS12, JS45 and AP-3 utilize the same fission pathway, encoded by similar genes, for the mineralization of this intermediate. This sequence and functional analysis of the aminophenol pathway have been studied in detail in AP3 (Park and Kim, 2001). It has a well-defined operonic structure that differs somewhat from the classic meta cleavage pathway, being specialized for the nitro substitutions. The presence of a similar aminophenol dioxygenase cleavage enzyme in Burkholderia xenovorans LB400 has led to the suggestion that the AP operon is widely distributed (Johnson and Spain, 2003). The regulatory module in the aminophenol operon is a MarR family repressor. A related gene cbaR was also found upstream of the chlorobenzoate catabolic genes in Tn5271, originally described in Comamonas testosteroni BR60, where it modulates induction of the catabolic genes (Providenti and Wyndham, 2001). The pathways being assembled for nitrobenzene therefore resemble those for we see for chlorinated aromatics: a widespread, but specialized ring-fission pathway is augmented by genes for the initial degradation, which are taken from different sources.

3.1.3

Atrazine

Degraders of the extensively used herbicide atrazine have recruited various genes for the same pathway from different sources, but the genes are often found in different locations on the genomes of the evolving degraders. The best known of the atrazine degradative plasmids is pADP1, a 108 kb plasmid found in Pseudomonas ADP, isolated from contaminated soil found at an agricultural chemical plant in Minnesota (Martinez et al., 2001). pADP-1 has an IncP-1b backbone 80–100% similar to that of R751 and other catabolic or multi-drug resistance plasmids of the IncP-1b group (Martinez et al., 2001; Schlu¨ter et al., 2007). It carries genes for the degradation of atrazine in two clusters. The enzymes required for initial steps, dechlorination, deamination and hydrolysis are encoded by atzA, atzB and atzC respectively. These seem to be recent arrivals on the plasmid backbone. They are expressed constitutively and are found dispersed (8–34 kb apart) between the tra and trb operons of the plasmid backbone. They are flanked by transposase genes (related to tnpA of IS801), and within the region lie two more copies of the TnpA, three

Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

29

copies of IS1071 and a mer operon (99% similar to that of Xanthomonas sp. W17). The atzA, atzB and atzC genes have been found together on different plasmids in phylogenetically diverse microorganisms: Chelatobacter neintzii, Stenotrophomonas maltophilia and Pseudoaminobacter (Topp et al., 2000b), and also in Agrobacterium, Clavibacter, Rhizobium, Pseudoalcaligenes, Alcaligenes and Ralstonia (Sajjaphan et al., 2004). They are also present in members of atrazine degrading consortia (De Souza et al., 1998a, b). The atzA gene appears to be more independent, or the most recently acquired gene – it is missing from some ADP variants and sometimes found alone in wild type bacteria (Bouquard et al., 1997). In contrast, genes for the degradation of the atrazine degradation intermediate cyanuric acid (produced via action of atzC) are regulated as one operon; atzD, atzE and atzF are located on the other side of the plasmid from atzA, atzB and atzC and are co-transcribed under the control of a lysR regulator. The DEF genes code for a cyanuric acid amidohydrolase, a nicotinamidase/pyrazinamidase and a urea amidolyase, which degrade cyanuric acid to biuret, to allphanate, and finally to ammonia and carbon dioxide. The atzE and atzF genes show 37 and 44% similarity to genes in Mycobacterium smegmatis and Saccharomyces cerevisiae, respectively (Martinez et al., 2001). Although the atzDEF genes are regulated as a unit, in contrast to the atzA, -B and -C genes, they too may have been assembled from diverse sources. Available evidence suggests that Gram-positive herbicide degrading bacteria use additional genes, or mix them with those of the Gram-negatives. trzN encodes a general triazine hydrolase and was originally described from a Nocardiodes strain (C190) (Mulbry et al., 2002; Topp et al., 2000a), which can degrade a range of chloro- and methylthio-substituted triazines. C190 does not have copies of known atzB or atzC genes. However, Arthrobacter crystallopoetes possesses trzN, atzB and atzC, as does Arthrobacter aurescens strain TC1. Strain TC1 contains a 380 kb plasmid pAA1 that has significant homology to pADP1 from Pseudomonas, and to pAO1 from Arthrobacter nicotinovorans (Sajjaphan et al., 2004) but pAO1 does not carry atz genes. However, A. nicotinovorans HIM can degrade atrazine and a number of related herbicides, and carries the atzABC genes on a 96 kb plasmid (Aislabie et al., 2005). Atrazine, a man-made compound, represents an example where degradation pathways must have been recently assembled from genes taken from both Gram negative and positive organisms. The recruitment of these genes has clearly been facilitated by broad-host-range plasmids, and by IS1071.

3.1.4

TSA – Toluenesulfonate

The degradative pathway for the detergent toluenesulfonate appears more assembled than those for atrazine and nitrobenzene in that most of the genes are found together on one element. In Comamonas testosteroni sp. T-2, initial steps are encoded by genes located together on the IncP-1b plasmid pTSA, in fact two copies are carried in a region flanked by two IS1071 (Tralau et al., 2001). Genes for a later step, encoded by psbA(C) are located on a separate plasmid, pT2L in this same strain T-2. The putative TSA transposon, carrying the initial steps, is found in TSA degraders from around the world, and is also found independently of IncP-1 plasmid backbones (Tralau et al., 2001). There are several TSA degraders that do not carry genes identical to those of T-2, so other variants of these genes, or different pathways exist. Nonetheless, the acquisition of TSA degradative abilities by bacteria is no doubt enhanced by the high mobility of the pTSA and the IS1071 elements.

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Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

Nylon Oligomers

Oligomers important to the synthesis of nylon are contaminants of nylon industry wastewaters, and a number of strains with the ability to degrade these oligomers have been isolated (Okada et al., 1983). Arthrobacter sp. strain K172 (previously classified as Flavobacterium) carries pOAD2 (Inc group not known; Negoro et al., 1983). The key genes nylA, nylB and nylC are all found on this plasmid, but they are scattered around, with one copy of nylB adjacent to nylC, a separately located copy of nylB, and also a separate nylA. Plasmid pOAD2 also contains five copies of IS6100 (Kato et al., 1994). The genes nylA and nylB were also found on separate plasmids (pNAD2 and pNAD6, respectively) in Pseudomonas sp. NK87. Plasmid pNAD2 also carries a copy of IS6100 (Kanagawa et al., 1989). This IS element is widely found associated with mobile elements of both Gram-positive and Gram-negative degraders, suggesting it plays an important role in assembly of this and other catabolic pathways. The nylB genes are highly homologous to chromosomal genes in both Arthrobacter aurescens TC1 (atrazine degrader) and Bradyrhizobium ORS5278. Work in this area seemed to halt, until the recent publication of the finding of variants of the nylC gene in Agromyces and Kocuria strains, isolated from cultures enriched at pH 10 (nylon oligomers are discharged from factories with alkaline solutions). nylA, B and C were found, the study focused on the thermal and pH stability properties of the NylC enzymes from both strains. They differed from the original by 5–15 amino acid residues (Yasuhira et al., 2007). Therefore, several version and vehicles of the nyl genes seem to exist, and they are highly transferable to wide range of species.

3.1.6

2,4-Dichlorophenoxacetic Acid

The degradation of the widely used herbicide 2,4-dichlorophenoxyacetic acid (2,4-D) has been intensely studied, and is a good example of pathway assembly through channeling of compounds into the modified ortho pathway. Its degradation is generally carried out by the action of the tfdA genes, encoding alpha-ketoglutarate dioxygenation, tfdB genes for oxygenation of the chlorophenol, and the widely conserved genes tfdCDEF for cleavage of the chlorocatechol. The tfdCDEF genes are also used by ortho cleavage utilizing chlorobenzoate degraders and are found in a growing list of proteobacteria. Most notably they are found in the multifunctional Burkholderia xenovorans LB400 and in Pseudomonas putida B13. An interesting genomic Island from the latter has been described (Gaillard et al., 2006). All of these were first found together, in duplicate, on the IncP-1b plasmid pJP4 (Laemmli et al., 2000; Perkins et al., 1990). The degradative region includes three copies of IS1071. This plasmid and others with rearranged key genes, are globally distributed. However, even on pJP4, the genes are not assembled into a single operon; some tfd-like genes are frequently found alone (i.e. tfdA found in many non 2,4-D degrading strains), and multiple homologues exist that are mixed in various strains and on different plasmid backbones (Fulthorpe et al., 1995; Top et al., 1995; Trefault et al., 2004; Vedler et al., 2004). Thus the 2,4-D pathway appears to be well assembled, but evidence for the independent origin and movement of the necessary genes is strong.

Evolution of New Catabolic Functions Through Gene Assembly by Mobile Genetic Elements

3.1.7

29

Carbazole

The 199 kb Inc P-7 plasmid pCAR1 from Pseudomonas resinovorans CA10 is another example of extensive mosaicity (Maeda et al., 2003). It contains the car and ant operons on a transposon, Tn4676, which encode the carbazole/dioxin-degrading enzymes and anthranilate 1,2-dioxygenase, respectively. According to the authors, the presence of remnants of numerous mobile genetic elements, as well as regions of higher and lower G + C content in the ant and car operon respectively, and the detection of carbazole degrading strains that carry Tn4657 or large contiguous fragments derived from Tn4657 in their chromosomes (Shintani et al., 2003, 2005), together suggest that assembly of Tn4676 may have occurred in several steps before it was captured in pCAR1. Inoue et al. (2004) reported virtually identical car genes in several carbazole degrading Pseudomonas sp. K23, and Janthinobacterium sp. J3. In these strains the operons are flanked by IS elements that differ from those found on pCAR1. More divergent but still related car genes have been recruited by carbazole degrading Sphingomonas strains. A plasmid of unknown incompatibility group, pCAR3, has been isolated from one of those strains, KA1 and fully sequenced – it has multiple carbazole degradation genes dispersed on four loci (Shintani et al., 2007; Urata et al., 2006). The carbazole example illustrates a complex case of widespread genes recruited by plasmids from different incompatibility groups in phylogenetically distinct degraders. Carbazole occurs naturally in tars and shale oils, so it is not surprising that the genes have been picked up by a wide variety of MGE. A more detailed analysis of the evolution of plasmid pCAR1 and its resemblance in structure to the TOL plasmid pWWO (not discussed here), has been described previously (Williams et al., 2004).

4

Research Needs

In conclusion, these different examples of retrospective evidence of catabolic pathway evolution in bacteria have given us tremendous insights, but many questions remain unanswered: Where, when and how frequently do these HGT steps take place, and how often do they result in a successfully assembled new pathway in a single organism? Compounds such as atrazine (De Souza et al., 1998a) and nitrotoluene (Snellinx et al., 2003) can either be degraded by a consortium of strains or by a single organism. But is the assembled pathway in one host really a few evolutionary steps ahead of the consortium, or do some environmental conditions and community structure promote multi-strain degradation while others select for the single degrading strain? Of intriguing interest is the observation that chlorobenzoate, a xenobiotic chemical, can induce the genes that facilitate genetic rearrangement (Sentchilo et al., 2003). What are the phylogenetic limits to catabolic gene recruitment? Are catabolic elements for xenobiotics really restricted to IncP1 elements? We need to keep in mind that the bacterial strains described above are among those that are easily grown in the laboratory, and several have been isolated after enrichment, introducing a great bias in the diversity of pollutant degrading bacteria we have studied so far. Thus, to further improve our insight in the evolution of degradation pathways in bacterial communities, we need comparative genomic studies of large numbers of plasmids, obtained through cultivation-independent studies, such as plasmid capture (Stuart-Keil et al., 1998; Top et al., 1995) and metagenomics (Ono et al., 2007), as well

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as from strains isolated using more diverse cultivation techniques (Romine et al., 1999). This retrospective evidence also needs to be complimented with direct detection of pathway assembly steps. Ideally we should be able to directly monitor gene transfer and rearrangement processes in microbial communities.

Acknowledgments This work was supported by an NSERC Discovery Grant to RRF and by NIH NIGMS grant R01GM073821 and NIH NCRR COBRE grant P20RR16448 to EMT. We thank M. Sota and H. Yano for valuable input.

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Fulthorpe RR, Mcgowan C, Maltseva OV, Holben WE, Tiedje JM (1995) 2,4-dichlorophenoxyacteic aciddegrading bacteria contain mosaics of catabolic genes. Appl Environ Microbiol 61: 3274–3281. Gaillard M, Vallaeys T, Vorholter F, Minola M, Werlen C, Sentchilo V, Puhler A, van der Meer J (2006) The clc element of Pseudomonas sp. strain B13, a genomic island with various catabolic properties. J Bacteriol 5: 1999–2013. Hacker J, Carniel E (2001) Ecological fitness, genomic islands and bacterial pathogenicity. EMBO Reports 21: 376–381. Inoue K, Widada J, Nakai S, Endoh T, Urata M, Ashikawa Y, Shintani M, Saiki Y, Yoshida T, Habe H, Omori T, Nojiri H (2004) Divergent structures of carbazole degradative car operons isolated from gramnegative bacteria. Biosci Biotechnol Biochem 68: 1467–1480. Jimenez JI, Minambres B, Garcia JL, Diaz E (2004) Genomic insights in the metabolism of aromatic compounds in Pseudomonas. In The Pseudomonads, vol. 3. Biosynthesis of Macromolecules and Molecular Metabolism. JL Ramos (ed.). New York: Kluwer, pp. 425–462. Johnson GR, Spain JC (2003) Evolution and catabolic pathways for synthetic compounds: bacterial pathways for degradation of 2,4-dinitrotoluene and nitrobenzene. Appl Microbiol Biotechnol 62: 110–123. Kanagawa K, Negoro S, Takada N, Okada H (1989) Plasmid dependence of Pseudomonas sp. strain NK87 enzymes that degrade 6-aminohexanoatecyclic dimer. J Bacteriol 171: 3181–3186. Kato K, Ohtsuki K, Mitsuda H, Yomo T, Negoro S, Urabe I (1994) Insertion sequence IS6100 on plasmid pOAD2, which degrades nylons oligomers. J Bacteriol 176: 11997–11200. Laemmli CM, Leveau JHJ, Zehnder AJB, van der Meer JR (2000) Characterization of a second tfd gene

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Ono A, Miyazaki R, Sota M, Ohtsubo Y, Nagata Y, Tsuda M (2007) Isolation and characterization of naphthalene-catabolic genes and plasmids from oil-contaminated soil by using two cultivationindependent approaches. Appl Microbiol Biotechnol 74: 501–510. Park H-S, Kim H-S (2000) Identification and characterization of the nitrobenzene catabolic plasmids pNB1 and pNB2 in Pseudomonas putida HS12. J Bacteriol 182: 573–580. Park H-S, Kim H-S (2001) Genetic and structural organization of the aminophenol catabolic operon and its implication for evolutionary process. J Bacteriol 183: 5074–5081. Perkins EJ, Gordon MP, Caceres O, Lurquin PF (1990) Organization and sequence analysis of the 2,4-dichlorophenol hydroxylase and dichlorocatechol oxidative operons of plasmid pJP4. J Bacteriol 5: 2351–2359. Pieper D, Reineke W (2004) Degradation of chloroaromatics by pseudomona(d)s. In The Pseudomonads, vol. 3. Biosynthesis of Macromolecules and Molecular Metabolism. JL Ramos (ed.). New York: Kluwer, pp. 509–574. Providenti MA, Wyndham RC (2001) Identification and functional characterization of CbaR, a MarR- Like modulator of the cbaABC-encoded chlorobenzoate catabolism pathway. Appl Environ Microb 67: 3530–3541. Ravatn R, Studer S, Springael D, Zehnder a JB, Van Der Meer JR (1998) Chromosomal integration, tandem amplification, and deamplification in Pseudomonas putida F1 of a 105-kilobase genetic element containing the chlorocatechol degradative genes from Pseudomonas sp. B13. J Bacteriol 180: 4360–4369. Reineke W (1998) Development of hybrid strains for the mineralization of chloroaromatics by patchwork assembly. Annu Rev Microbiol 52: 287–331. Reineke W, Knackmuss H-J (1988) Microbial degradation of haloaromatics. Ann Rev Microbiol 42: 263–287. Romine MF, Stillwell LC, Wong K-K, Thurston SJ, Sisk EC, Sensen C, Gaasterland T, Fredrickson JK, Saffer JD (1999) Complete sequence of a 184kilobase catabolic plasmid from Sphingomonas aromaticivorans F199. J Bacteriol 181: 1585–1602. Rousseaux S, Soulas G, Hartmann A (2002) Plasmid localisation of atrazine-degrading genes in newly described Chelatobacter and Arthrobacter strains. FEMS Microbiol Ecol 41: 69–75. Sajjaphan K, Shapir N, Wackett LP, Palmer M, Blackmon B, Tomkins J, Sadowsky MJ (2004) Arthrobacter aurescens TC1 atrazine catabolism genes trzN, atzB, and atzC are linked on a 160-kilobase region and are functional in Escherichia coli. Appl Environ Microbiol 70: 4402–4407.

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Top EM, Springael D (2003) The role of mobile genetic elements in bacterial adaptation to xenobiotic organic compounds. Curr Opin Biotechnol 14: 262–269. Top EM, Holben WE, Forney LJ (1995) Characterization of diverse 2,4-dichlorophenoxyacetic acid-degradative plasmids isolated from soil by complementation. Appl Environ Microbiol 61: 1691–1698. Top EM, Moenne-Loccoz Y, Pembroke T, Thomas CM (2000) Phenotypic traits conferred by plasmids. In The Horizontal Gene Pool. CM Thomas (ed.). Amsterdam: Harwood Academic, pp. 249–286. Topp E, Mulbry WM, Zhu H, Nour SM, Cuppels D (2000a) Characterization of S-triazine herbicide metabolism by a Nocardioides sp. isolated from agricultural soils. Appl Environ Microbiol 66: 3134–3141. Topp E, Zhu H, Nour SM, Houot S, Lewis M, Cuppels D (2000b) Characterization of an atrazine-degrading Pseudaminobacter sp. isolated from Canadian and French agricultural soils. Appl Environ Microbiol 66: 2773–2782. Toussaint A, Merlin C, Monchy C, Benotmane M, Leplae R, Mergeay M, Springael D (2003) The biphenyland 4-chlorobiphenyl-catabolic transposon Tn4371, a member of a new family of genomic islands related to IncP and Ti plasmids. Appl Environ Microbiol 69: 4837–4845. Tralau T, Cook a M, Ruff J (2001) Map of the IncPbeta plasmid pTSA encoding widespread genes (tsa) for p-toluenesulfonate degradation in Comamonas testosteroni T-2. Appl Environ Microbiol 67: 1508–1516. Trefault N, De La Iglesia R, Molina a M, Manzano M, Ledger T, Perez-Pantoja D, Sanchez MA, Stuardo M, Gonzalez B (2004) Genetic organization of the catabolic plasmid pJP4 from Ralstonia eutropha JMP134 (pJP4) reveals mechanisms of adaptation to chloroaromatic pollutants and evolution of specialized chloroaromatic degradation pathways. Environ Microbiol 6: 655–668. Tsuda M, Tan HM, Nishi A, Furukawa K (1999) Mobile catabolic genes in bacteria. J Biosci Bioeng 87: 401–410. Urata M, Uchimura H, Noguchi H, Sakaguchi T, Takemura T, Eto K, Habe H, Omori T, Yamane H, Nojiri H (2006) Plasmid pCAR3 contains multiple gene sets involved in the conversion of carbazole to anthranilate. Appl Environ Microbiol 72: 3198–3205. van der Meer JR (2007) A genomic view on the evolution of catabolic pathways and bacterial adaptation to toxic compounds. In: Microbial Biodegradation. Genomics and Molecular Biology E Diaz (ed.). Norfolk UK: Caister Academic Press, pp. 219–269. van der Meer, JR, de Vos WM, Harayama S, Zehnder AJ (1992) Molecular mechanisms of genetic adaptation

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30 Experimental Evolution of Novel Regulatory Activities in Response to Hydrocarbons and Related Chemicals V. Shingler Department of Molecular Biology, Umea˚ University, Umea˚, Sweden [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1236 2 Applications of Nature’s Sensor-Regulators . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1236 3 Methodology for Generating Novel Response Profiles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1238 4 Screening and Selecting Sensor-Regulators with Desirable Properties . . . . . . . . . . . . . 1239 5 Limitations Imposed by Effector-Responses of Sensor Regulators . . . . . . . . . . . . . . . . . . 1239 6 Mechanistic Interpretation of Effector-Response Mutants . . . . . . . . . . . . . . . . . . . . . . . . . . 1240 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1243

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_86, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Bacterial regulatory proteins that control catabolism of hydrocarbons and related chemicals have evolved (or are actively evolving) towards specifically detecting compounds that signal the presence of growth substrates. Laboratory evolution of the chemical-binding and response properties of sensory-regulators has been achieved by a number of different techniques to generate novel derivatives with desired properties. Such manipulated and selected regulatory proteins are increasingly used in artificial genetic circuitry for improved biodegradation systems, biosensor construction, and in assembling regulatory cascades for synthetic biology within a wide range of biotechnological applications.

1

Introduction

Regulators that control gene expression are nature’s sensing systems for detection of substrates, chemicals and metabolites. The sensor-regulatory proteins that govern catabolism of hydrocarbons and related compounds include representatives from all major families of bacterial regulators and function by the general molecular mechanisms of their given class to control transcription. With the exception of two component systems that are comprised of separate sensory and regulator proteins, the chemical-sensing property is directly built into a transcriptional regulator so that binding of specific substrates or intermediates of the pathway it controls directly alters its regulatory activity (Shingler, 2003). Thus, sensor-regulators that directly couple chemical-detection with transcriptional performance represent an immense resource for the generation of improved or optimized regulatory systems. In principle, the binding and/or response properties of any sensory-regulator can be manipulated to either expand or contract the range of compounds that affects its transcriptional promoting properties, i.e., its effector specificity profile. In this essay I draw attention to selected studies that illustrate how the effector specificity of single component sensorregulators can be artificially evolved, recombined, mutated or otherwise modified to create regulators with desired alterations in their response profiles towards hydrocarbons and related compounds. However, the experimental strategies, applications and concepts outlined below could also be readily applied to manipulations of two component regulatory systems.

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Applications of Nature’s Sensor-Regulators

Sensory-regulators with appropriate chemical-binding properties can be harnessed in a number of biotechnological applications (> Fig. 1). Prevalent among these are their use as the biological detection component of whole cell biosensors for monitoring available, and thus biologically relevant, levels of compounds in environmental samples (> Fig. 1b, van der Meer et al., 2004). The chemical-binding properties of sensor-regulators are also used to provide appropriate regulatory circuits for engineered catabolic pathways and for artificial regulatory cascades (> Fig. 1b, Cebolla et al., 2001; de Las Heras et al., 2008; Royo et al., 2007; Silva-Rocha and de Lorenzo, 2008), and are increasingly employed to create genetic traps to identify genes encoding new catabolic enzymes (> Fig. 1c, Mohn et al., 2006; Uchiyama et al., 2005; van Sint Fiet et al., 2006). These applications of sensory-regulators require defined windows of activity that span only desired responses. Such desirable responses frequently include novel

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. Figure 1 Examples of biotechnological applications of sensor-regulators that respond to hydrocarbons and related compounds. (a) In their natural context, sensor-regulators (S-R), be they repressors or activators, are usually encoded next to the genes they control. Binding of the effector compound (E) triggers transcription (via derepression or activation) from the promoter (P), leading to expression of the downstream genes only in the presence of the effector. (b) Similar systems are adopted to control (1) synthetically constructed pathway operons and (2) biosensors, which in their simplest form appropriate a suitable chemical-responsive transcriptional regulator and the cognate promoter it controls and couples the regulatory circuit to the output of a readily detectable gene product (e.g., color [b-galactosidase], light [luciferase, Lux, Luc], or fluorescence [gfp]). Alternatively, the adopted regulatory circuit can be coupled to control the expression of a second regulatory protein (3) to amplify the original signal provided by the presence of a compound. (c) Sensor-regulator circuits can also be adopted for metabolic mining. Under this scenario, a biosensor system (2) is coincorporated with a genetic system that allows expression of genomic or meta-genomic DNA that potentially encodes for biotransformation of a compound (X) to an effector of the sensor-regulator. Thus, a positive transcriptional read-out only occurs if a gene encoding the desired enzymatic activity has been productively cloned and expressed.

or improved ability to promote transcription in response to man-made or recalcitrant compounds (Galvao and de Lorenzo 2006). Conversely, undesirable responses to structurally related compounds often need to be suppressed, because they reduce the specificity of biosensors or enzymatic traps and would lead to inappropriate and potentially deleterious

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expression of catabolic enzymes. Thus, strategies for selection and characterization of regulators with optimal sensory-response properties are integral to biotechnological and synthetic biology development.

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Methodology for Generating Novel Response Profiles

Many possible approaches have been, or could be, used to generate genetic diversity to modify or generate new effector response profiles (> Fig. 2). Random (blind) mutagenesis approaches target the entire coding region or subregions for domains known to be involved in effectorbinding or responses and can be directly applied even if structural or mechanistic information is lacking. Such blind strategies include isolation of spontaneous mutants (e.g., DmpR, Sarand et al., 2001), those produced through chemical mutagenesis (e.g., XylS and XylR, Delgado and Ramos 1994, Michan et al., 1992), error-prone PCR (e.g., DmpR, HbpR and NahR, Cebolla et al., 1997; Wise and Kuske, 2000; Beggah et al., 2008) or DNA shuffling (e.g., DmpR and XylR, Ska¨rfstad et al., 2000). The availability of mechanistic or structural models of sensor-regulators has also opened up the possibility of site-specific mutagenesis of effector-binding sites to create novel binding capacities and more specific or sensitive regulators (e.g., NahR and DntR, Park et al., 2005; Lonneborg et al., 2007). An alternative strategy is to use computational modeling to design specific binding sites within protein scaffolds that are not natively responsive to hydrocarbons and related compounds. This is exemplified by the introduction of ligand docking sites for nitrotoluenes and trinitrobenzene within periplasmic binding proteins for biosensor applications (Looger et al., 2003).

. Figure 2 Strategic steps for generating, identifying and refining the activities of desirable mutant sensorregulators.

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Screening and Selecting Sensor-Regulators with Desirable Properties

Irrespective of the strategy used to obtain a pool of potential variants, screening or selection strategies need to be applied to identify those that positively respond to the target compound (> Fig. 2). Most screening procedures are conceptually similar to biosensors (> Fig. 1b2), in which the activities of variants are monitored through the activity of reporter genes (e.g., b-galactosidase, luciferases or fluorescent proteins) in response to potential effectors in the growth medium. In the later case, identification of derivatives with new response properties can be made high throughput using fluorescence activated cell sorting (e.g., Beggah et al., 2008). As an alternative to screens, genetic selections avoid consideration of the vast majority of mutations that result in subresponsive or inactive variants. In such genetic selections, regulator-dependent promoters are coupled to selectable markers that mediate, for example, growth at the expense of a particular carbon source (Sarand et al., 2001), growth in the absence of a particular amino acid (Galvao et al., 2007), or in the presence of an antibiotic (Michan et al., 1992; Pavel et al., 1994). Both screens and selections suffer from the common feature of also readily identifying fully or partially constitutively active variants that do not require an effector to be able to activate transcription. The strategies outlined above make it relatively easy to identify variants that respond to the presence of a target compound. However, constricting or abolishing undesirable responses presents a more difficult task. Genetic systems that positively select for the absence of an undesirable response are few and far between. The sacB system, in which expression of the SacB protein causes lysis in the presence of sucrose, has been used to counter-select activities of DmpR and XylR (Garmendia et al., 2001; Ng et al., 1996). However, this genetic system, which operates well with Escherichia coli, is sometimes inefficient in other bacteria. The amiE gene that encodes an aliphatic amidase capable of converting fluoroacetamide to the toxic compound fluoroacetate has been successfully used in an analogous counter-selection regime in Pseudomonas aeruginosa (Collier et al., 2001). Similarly, the pyrF, a yeast URA3 homologue, associated with uracil prototrophy and flouroorotic acid sensitivity offers another potential solution to this problem (Galvao et al., 2007).

5

Limitations Imposed by Effector-Responses of Sensor Regulators

Native sensor-regulators vary extensively in the number and range of compounds they respond to (> Fig. 3). Some LysR-type regulators, for example, are responsive only to a specific intermediate generated through the catabolic pathway they control, while some members of the s54-dependent family of bacterial enhancer-binding proteins (bEBPs) have very broad effector response profiles that include initial pathway substrates, intermediates, as well as nonmetabolizable but structurally related analogues (reviewed in Shingler 2003). It has long been recognized that the response profiles of sensor-regulators can potentially limit catabolic performance through poor or nonresponsiveness to compounds that can be metabolized through cognate pathways. Experimental evolution of mutants that remove regulatory bottlenecks through acquiring novel or enhanced ability to respond to a target compound can enhance catabolic performance (e.g., Ramos et al., 1986; Pavel et al., 1994).

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. Figure 3 Functional domains of experimentally evolved sensor-regulators for hydrocarbons and related compounds. Specific members of the individual families discussed in the text are given in brackets. HTH stands for helix-turn-helix motif and indicates the involvement in DNA-binding. Sensory indicates regions identified as interacting with the effector compound, while activation indicates domains involved in promoting transcription by RNA polymerase bound to cognate promoters as detailed in the text. V4R (pfam02830.8) indicates the location (residues 119–196) of an ancient hydrocarbon binding signature motif found in the sensory domains of the ligand responsive subgroup of bEBPs that closely overlaps the major effector-specificity subregion of the sensory domain (residues 110–186) identified by DNA shuffling (Ska¨rfstad et al., 2000). GAFTGA/AAA+ (pfam00004.8) signature module is as described in the text.

Such plasticity in the effector-responses of sensor-regulators is not only observed in the laboratory. Poor recognition of 4-methylphenol by DmpR limits biodegradative capacity because it elicits suboptimal expression of the enzymes necessary for its catabolism (Pavel et al., 1994). Some DmpR mutants were isolated simply by recovering a P. putida strain carrying DmpR after 2–4 days exposure to soil amended with 4-methylphenol. This procedure led to the isolation of subpopulations with improved ability to degrade 4-methylphenol. Enhanced growth properties of individual isolates were a result of different spontaneous mutations within the DmpR effector-binding domain that alter the sensitivity and the range of effectors that elicited expression of the catabolic enzymes (Sarand et al., 2001). These results suggest that at least in this case, altered regulator effector-responses are an easy road to improved catabolic fitness that is likely to be a continual and ongoing adaptation mechanism to optimize catabolism.

6

Mechanistic Interpretation of Effector-Response Mutants

For repressor and activator sensor-regulators, effector-binding ultimately controls transmission of steps leading to transcriptional activation. While sensor-regulators are found within all major families of bacterial regulators, representatives of the LysR-, AraC/XylS- and bEBPs (> Fig. 3) have been most extensively studied with respect to mutants that alter effectorspecificity responses to hydrocarbons and related chemicals. (A) LysR-type transcriptional regulators Based on sequence homologies and/or domain prediction, LysR-type transcriptional regulators (LTTRs) are typically 300 amino acids (aa) in length and widely distributed in bacteria.

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Structures of a number of LTTRs are available and include CysB (truncated), Cbl (truncated), OxyR (truncated), CbnR, and those of wild-type and two effector-response mutants of DntR (Lonneborg et al., 2007 and references therein). As illustrated in > Fig. 3, common to all LTTRs is a 60–70 aa N-terminal helix-turn-helix (HTH) DNA binding domain. The DNA binding domain is coupled through a poorly conserved effector-binding sensory region (consisting of two subregions between  residues 95 and 173, and 196 and 206) to a C-terminal region, which is involved in tetramerization of two head-to-tail dimers and transcriptional activation (reviewed in Tropel and van der Meer 2004). Effector-binding to LTTRs is thought to trigger conformational changes that are channeled to the DNA-binding interface to facilitate interaction with RNA polymerase and subsequent open-complex formation (Tropel and van der Meer, 2004). NahR and DntR are two salicylate-responsive members of this family whose effector responses can really be expanded or converted to accommodate benzoate by either blind or site directed approaches. Both approaches have lead to the identification of some derivatives that can respond to 3-chlorobenzoate, 4-nitrobenzoate or 3-methyl salicylate (Cebolla et al., 1997; Lonneborg et al., 2007; Park et al., 2005). Both blind and site-directed approaches to manipulate the effector-responses of NahR and DntR have coincided in pinpointing residues 169 and 248 – that based on the DntR structure lie within the predicted effector-binding cavity – as key residues for effector specificity (Galvao and de Lorenzo 2006). However, more extensive mutagenesis of these residues (either alone or together) in NahR (Park et al., 2005), and additional residues within the predicted effector-binding cavity of DntR (Lonneborg et al., 2007) highlight that effector-binding affinity does not necessarily translate to efficient transcriptional activation. For example, combining mutations of residues F111 and H169 of DntR, which would each individually be predicted to decrease affinity for salicylate, resulted in higher transcription in response to salicylate (Lonneborg et al., 2007). Moreover, site specific mutations designed to accommodate liganding of 2,4-dinitrotoluene resulted in unexpected movement of other residues that likely resulted in detrimental effects in communicating the effector-binding signal to productive transcriptional activation. (B) AraC/XylS-type transcriptional regulators Of this large family of regulators, XylS is the most extensively characterized member that responds to hydrocarbons and related chemicals. Regulators of this protein family typically possess an approximately 100 amino acid C-terminal region containing two HTH motifs that are sufficient for DNA-binding and transcriptional activation, linked to a nonconserved N-terminal region implicated in effector-binding, dimerization and interaction with the transcriptional machinery (reviewed in Tropel and van der Meer 2004). Despite the availability of structural models for family members MarR and Rob (Rhee et al., 1998; Kwon et al., 2000), low sequence conservation within the N-terminal of the different family members has hampered identification of key residues involved in binding diverse effectors. For dimeric AraC, binding of the sugar arabinose mediates a conformational switch to the active form which involves simultaneous coverage of the effector-binding pocket and releases of the DNA binding domains to enabling transcriptional activation (Schleif, 2003). Analysis of derivatives of XylS in response to its native effector 3-methylbenzoate has provided evidence that its effector activation mechanism likewise involves derepression of inhibitory interdomain interactions to release its DNA binding determinants (Dominguez-Cuevas et al., 2008). Both blind and site directed mutagenesis approaches have been used to generate variants of XylS derivatives with expanded effector activation profiles that include benzoates with

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ethyl-, chloro-, methyl-, methoxy- and/or hydroxy-substitutions at positions that do not function in activating wild-type XylS (Michan et al., 1992; Ramos et al., 1986). These effector response mutants of XylS pin-pointed an 11 amino acid glycine-rich N-terminal patch that is likely to be involved in effector-recognition, although mutations that alter the effectorspecificity and dependence map throughout the protein. Thus, as with members of the LTTR family, these studies highlighted the importance of interdomain communication in coupling the effector-binding response to transcriptional activation. (C) s54-dependent family of bacterial enhancer-binding proteins Initiation of transcription from promoters recognized by holoenzyme RNA polymerase associated with the s54 subunit strictly requires activation by a mechano-regulator. Members of this family of transcriptional activators use ATP-catalysis to remodel s54-RNAP locked in the inactive closed complex to stimulate DNA melting and formation of the functional open complex. This activation process is generally mediated by the activator bound to enhancer sequences located approximately 100–200 base pairs from the promoter they control, thus lending the family the name bacterial enhancer binding proteins or bEBPs. The hallmark of bEBPs is an 300 aa AAA + homology domain for ATP-binding and hydrolysis that contains two mobile loops: the GAFTGA loop that communicates with s54-RNAP and a so called loop 2 that is believed to be involved in sensing and communicating the nucleotide state (reviewed in Schumacher et al., 2006). The GAFTGA/AAA + module is usually linked to a DNA binding domain at its C-terminus, and in most cases to a regulatory domain at its N-terminus (> Fig. 3), although members of this family lacking one or both of the latter two domains do exist (Beck et al., 2007). The distinct effector response profiles of a subgroup of bEBPs that respond to the presence of hydrocarbons and related compounds, are generally broad (reviewed in Galvao and de Lorenzo 2006). For this subgroup of bEBPs, which includes DmpR, HbpR and XylR, the N-terminal sensory-domain is sufficient to bind the effector compound and, at least in the case of DmpR, it has been demonstrated that a single site is used for the response to multiple different effectors (O’Neill et al., 1999). Direct binding of chemicals to the N-terminal sensory domain mediates a conformational change that releases repressive interactions of the sensory domain on the AAA + module and thereby allows ATP-binding triggered multimerization to the active form (Wikstro¨m et al., 2001 and references therein). A small structured linker that joins the sensory A-domain to the AAA + module in DmpR, XylR, and many other hydrocarbon-responsive members, is intimately involved in coordinating this effector-binding response (O’Neill et al., 2001). Thus, both effector-binding and a concomitant switching to a multimeric state that is capable of correct interfacing with s54RNAP are required for a productive transcriptional response from bEBPs. DNA shuffling between DmpR and XylR identified a subregion of the N-terminal domain ( residues 110–186) that contains the major effector-specificity determinants and which encompasses an apparently ancient hydrocarbon-binding signature motif (V4R) found in proteins from all domains of life (> Fig. 3). Random selection of variants of this subgroup has identified many structural variants with substitutions within this region which result in novel response properties. Interestingly, however, acquisition of novel response properties does not necessarily involve creation of a new effector-binding property, rather where tested it appears to lie within the capability of a preexisting ability to bind the effector to productively couple to a conformation capable of promoting transcription (Garmendia et al., 2001; O’Neill et al., 1999). While many substitutions that alter the effector response profiles of

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DmpR and XylR do map within this region, others do not, a property that is particularly prevalent when isolating XylR derivatives able to respond to 2,4-dinitrotoluene (Galvao et al., 2007). A robust bioinformatics-based model for XylR that can explain the phenotypes of many DmpR and XylR mutations has been generated. Though it remains to be experimentally tested, this model suggests that it is not only the geometry of the binding site per se, but also a conduit made up from surfaces of the N-terminal sensory domain and AAA + modules that determines the response to effector-binding (Galvao and de Lorenzo 2006 and references therein).

7

Research Needs

Computational design offers enormous potential for engineering ligand docking sites (Galvao and de Lorenzo 2006). The caveat is that the systematic manipulation of binding sites for hydrocarbons and related compounds to generate predictable responses still remains a major challenge. Both blind and directed experimental evolution as strategies for engineering novel recognition and response properties of sensor regulators have undoubtedly both increased mechanistic understanding of different regulatory classes of these types of proteins and generated derivatives with novel response properties towards hydrocarbons and related compounds that have potential utility within biotechnological applications. However, as emphasized here, the series of events that communicates the effector-binding signal to unlock the transcriptional regulatory properties is still poorly understood, and mutations that alter effector-binding affinity can have surprising and unpredictable effects on transcriptional output. Thus, for many sensor-regulators a greater understanding of interdomain communication is required before computational-design based alterations in ligand binding can be translated to fully predictable outcomes. A reoccurring theme in blind screens and genetic selections is the identification of mutant regulators that are altered in positions away from a proposed binding site that could not have been predicted on the basis of current structural and/or mechanistic information. Thus, in the absence of a full understanding of transmission pathways for effector-binding, blind approaches combined with robust screens and/or genetic selections appear to remain the most potent and shortest route to isolate derivatives with desired properties. The attainment of specificity for a novel effector requires artificial sequential rounds of mutagenesis and activity refinement to remove concomitant undesirable responses to other effectors (> Fig. 2). In this respect, a greater array of techniques to counter select undesirable properties is needed for appropriate activity refinement for applications that require unique and/or limited response profiles. Another bias to overcome is the use of known regulatory systems. Harvesting and modifying genetic information from the metagenome provides an enormous resource of potential regulators with the ability to bind and respond to different target compounds. However, as discussed elsewhere (Cases and de Lorenzo 2005; Shingler 2003), sensorregulators and the promoters they control do not function as isolated units, rather they are nested within the evolutionary selected global regulatory networks of host cells. These global regulatory systems can greatly influence the performance of a given regulatory circuit dependent on its genome context and copy number. Hence, selections of sensor-regulators with a particular desired property would most effectively be performed within the genetic context in which they are destined to operate.

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Acknowledgments Research in the Shingler laboratory is supported by the Swedish Research Council.

References Beck LL, Smith TG, Hoover TR (2007) Look, no hands! Unconventional transcriptional activators in bacteria. Trends Microbiol 15: 530–537. Beggah S, Vogne C, Zenaro E., van de Meer JR (2008) Mutant HbpR transcription activator isolation for 2-chlorobiphenyl via green flourescent proteinbased flow cytometry and cell sorting. Microb Biotechnol 1: 68–78. Cases I, de Lorenzo V (2005) Promoters in the environment: transcriptional regulation in its natural context. Nat Rev Microbiol 3: 105–118. Cebolla A, Sousa C, de Lorenzo V (1997) Effector specificity mutants of the transcriptional activator NahR of naphthalene degrading Pseudomonas define protein sites involved in binding of aromatic inducers. J Biol Chem 272: 3986–3992. Cebolla A, Sousa, C, de Lorenzo V (2001) Rational design of a bacterial transcriptional cascade for amplifying gene expression capacity. Nucleic Acids Res 29: 759–766. Collier DN, Spence C, Cox MJ, Phibbs PV (2001) Isolation and phenotypic characterization of Pseudomonas aeruginosa pseudorevertants containing suppressors of the catabolite repression control-defective crc-10 allele. FEMS Microbiol Lett 196: 87–92. de Las Heras A, Carreno CA, de Lorenzo V (2008) Stable implantation of orthogonal sensor circuits in Gramnegative bacteria for environmental release. Environ Microbiol 10: 3305–3316. Delgado A, Ramos JL (1994) Genetic evidence for activation of the positive transcriptional regulator Xy1R, a member of the NtrC family of regulators, by effector binding. J Biol Chem 269: 8059–8062. Dominguez-Cuevas P, Marin P, Busby S, Ramos JL, Marques S (2008) Roles of effectors in XylSdependent transcription activation: intramolecular domain derepression and DNA binding. J Bacteriol 190: 3118–3128. Galvao TC, de Lorenzo V (2006) Transcriptional regulators a la carte: engineering new effector specificities in bacterial regulatory proteins. Curr Opin Biotechnol 17: 34–42. Galvao TC, Mencia M, de Lorenzo V (2007) Emergence of novel functions in transcriptional regulators by regression to stem protein types. Mol Microbiol 65: 907–919.

Garmendia J, Devos D, Valencia A, de Lorenzo V (2001) A la carte transcriptional regulators: unlocking responses of the prokaryotic enhancer-binding protein XylR to non-natural effectors. Mol Microbiol 42: 47–59. Kwon HJ, Bennik MH, Demple B, Ellenberger T (2000) Crystal structure of the Escherichia coli Rob transcription factor in complex with DNA. Nat Struct Biol 7: 424–430. Lonneborg R, Smirnova I, Dian C, Leonard GA, Brzezinski P (2007) In vivo and in vitro investigation of transcriptional regulation by DntR. J Mol Biol 372: 571–582. Looger LL, Dwyer MA, Smith JJ, Hellinga HW (2003) Computational design of receptor and sensor proteins with novel functions. Nature 423: 185–190. Michan C, Zhou L, Gallegos MT, Timmis KN, Ramos JL (1992) Identification of critical amino-terminal regions of XylS. The positive regulator encoded by the TOL plasmid. J Biol Chem 267: 22897–22901. Mohn WW, Garmendia J, Galvao TC, de Lorenzo V (2006) Surveying biotransformations with a la carte genetic traps: translating dehydrochlorination of lindane (gamma-hexachlorocyclohexane) into lacZ-based phenotypes. Environ Microbiol 8: 546–555. Ng LC, O’Neill E, Shingler V (1996) Genetic evidence for interdomain regulation of the phenol-responsive final s54-dependent activator DmpR. J Biol Chem 271: 17281–17286. O’Neill E, Sze CC, Shingler V (1999) Novel effector control through modulation of a preexisting binding site of the aromatic-responsive s54dependent regulator DmpR. J Biol Chem 274: 32425–32432. O’Neill E, Wikstrom P, Shingler V (2001) An active role for a structured B-linker in effector control of the s54-dependent regulator DmpR. EMBO J 20: 819–827. Park HH, Lee HY, Lim WK, Shin HJ (2005) NahR: effects of replacements at Asn 169 and Arg 248 on promoter binding and inducer recognition. Arch Biochem Biophys 434: 67–74. Pavel H, Forsman M, Shingler V (1994) An aromatic effector specificity mutant of the transcriptional regulator DmpR overcomes the growth constraints of Pseudomonas sp. strain CF600 on

Experimental Evolution of Novel Regulatory Activities para-substituted methylphenols. J Bacteriol 176: 7550–7557. Ramos JL, Stolz A, Reineke W, Timmis KN (1986) Altered effector specificities in regulators of gene expression: TOL plasmid xylS mutants and their use to engineer expansion of the range of aromatics degraded by bacteria. Proc Natl Acad Sci USA 83: 8467–8471. Rhee S, Martin RG, Rosner JL, Davies DR (1998) A novel DNA-binding motif in MarA: the first structure for an AraC family transcriptional activator. Proc Natl Acad Sci USA 95: 10413–10418. Royo JL, Becker PD, Camacho EM, Cebolla A, Link C, Santero E, Guzman CA (2007) In vivo gene regulation in Salmonella spp. by a salicylate-dependent control circuit. Nat Methods 4: 937–942. Sarand I, Skarfstad E, Forsman M, Romantschuk M, Shingler V (2001) Role of the DmpR-mediated regulatory circuit in bacterial biodegradation properties in methylphenol-amended soils. Appl Environ Microbiol 67: 162–171. Schleif R (2003) AraC protein: a love-hate relationship. Bioessays 25: 274–282. Schumacher, J, Joly, N, Rappas, M, Zhang, X, Buck, M (2006) Structures and organisation of AAA + enhancer binding proteins in transcriptional activation. J Struct Biol 156: 190–199. Shingler V (2003) Integrated regulation in response to aromatic compounds: from signal sensing to attractive behaviour. Environ Microbiol 5: 1226–1241.

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31 Rational Construction of Bacterial Strains with New/Improved Catabolic Capabilities for the Efficient Breakdown of Environmental Pollutants R.-M. Wittich . P. van Dillewijn . J.-L. Ramos* Department of Environmental Protection, Estacio´n Experimental del Zaidı´n, Consejo Superior de Investigaciones Cientı´ficas, Granada, Spain *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1248 2 Improved Catabolic Enzymes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1249 3 Metagenomic Strategies to Mine the Enzymes of Uncultivable Bacteria . . . . . . . . . . 1250 4 Improved Catabolic Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1250 5 Microbial Releases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1251 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1253

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_87, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Although numerous bacteria capable of degrading environmental pollutants have been isolated, characterized and used in bioremediation processes, many compounds of environmental concern such as PCBs, dioxins, and explosives are still hardly biodegradable. The emerging methodologies of genetic engineering, although considered as highly promising for the construction of ‘‘superbugs,’’ largely failed to create the requested recombinant hyperdegraders with catabolical enzymes exhibiting novel and/or highly improved features, apart from very few exceptions. The generation of these ‘‘superbugs’’ will require additional cellular evolution and a more profound understanding and fine-tuning of genetic regulation which could be helped by the exploitation of the metagenome and novel strategies in synthetic biology.

1

Introduction

Within the last 5 decades numerous bacterial strains have been isolated from environmental samples which are capable of degrading environmental pollutants such as polycyclic aromatic hydrocarbons (PAHs), halophenols, halobenzenes, haloaliphatics, nitro- and sulfoaromatics, and many pesticides. Investigation of the taxonomical, physiological, biochemical and genetic characteristics of these strains led to the establishment of economically important biotechniques for environmental cleanup. Bio-augmentation and biostimulation became catchwords in applied science and bioindustries. On the other hand, the elucidation of catabolic pathways for xenobiotic degradation/transformation and the availability of a large number of genetic methodologies which allow the rational design of genes with predetermined characteristics led to the quest for creating bacterial cells (‘‘superbugs’’) which could degrade all compounds of environmental concern. However, little has been published until recently in scientific journals on really new constructs exhibiting significantly better biodegradation capacities and survival in contaminated environments. Moreover, since the late 1970s there has been a dearth (lack) in discoveries of new biochemical reactions with enzymes with truly novel functions. As a result and as discussed by Cases and de Lorenzo (2005), the strategies propagated since the mid 1980s for the construction of ‘‘superbugs’’ for bioremediation purposes have largely failed to date. Nevertheless, more efficient bacteria are still needed for the degradation of complex (more than one single aromatic ring system) compounds such as PCBs, chlorinated dibenzofurans and -dioxins, polyhalogenated diphenyl ethers, pesticides like DDT and related structures; polynitroaromatics like TNT, etc. Here we discuss some ideas for the rational construction of bacterial strains with new/ improved catabolic capabilities for the efficient breakdown of environmental pollutants. In order to rationally construct effective bacteria which can degrade environmental pollutants a larger pool of effective catabolic enzymes are required. Enrichment of cultivable bacteria from polluted sites has revealed a finite number of catabolic pathways. Often ‘‘novel’’ enzymes associated with new isolates are described in literature but their novelty is usually due to phylogenetic differences with enzymes with similar activities rather than to novel functions. Moreover, often novel enzymes are not compared with their counterparts to determine which enzyme is more efficient in degrading a pollutant. Alternative strategies are being used and devised to (1) improve known enzyme activities and (2) to find truly novel biochemical pathways. On the other hand effective catabolic strains need to be found or developed to express improved catabolic pathways to efficiently degrade pollutants on the one hand and which

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can survive in the environment for sufficient time to remediate polluted sites. As this implies the release of genetically modified organisms (GMOs) this topic will be briefly addressed as well.

2

Improved Catabolic Enzymes

An effective tool used to improve degradative enzymes is by in vitro evolution technologies. These techniques have been used for PCB degradation by biphenyl dioxygenases to extend the substrate range and the regiospecificity of (oxygenolytic) attack on defined PCB congeners (see reviews by Furukawa, 2003; Furukawa et al., 2004; Kimbara, 2005; Pieper, 2005). Similarly, heme-containing monooxygenases such as P450CAM were modified and improved. The CYP101 monooxygenase activity towards the oxidative dehalogenation and/or hydroxylation of highly chlorinated benzenes allowed pentachlorobenzene to be transformed into pentachlorophenol and the same compound obtained from hexachlorobenzene at a reasonable rate of transformation. The same modified CYP101 could transform the less halogenated 1,3,5-trichlorobenzene, a structural isomer hitherto identified as recalcitrant towards bacterial oxidation and aerobic mineralization, into 2,4,6-trichlorophenol (Chen et al., 2002), a compound known to be mineralized by the 2,4-D degrader Cupriavidus necator JMP134 (Clement et al., 1995). On the other hand catabolic bacterial processes can be engineered to combine oxic and anoxic enzymatic reactions. For a long time P450CAM monooxygenases have been known to catalyze dehalogenations of aliphatic compounds. These enzymes were subject to alterations because these reactions did not necessarily require oxygen. The papers of Wackett et al. (1994) and Hur et al. (1994) demonstrated for the first time that such an engineered heme oxidase can be used to reductively dehalogenate polyhalogenated alkanes to haloalkenes and the latter oxidatively dehalogenated by a genetically optimized dioxygenase to glyoxylate and even CO2 when these activities were combined in an appropriate bacterial host such as Pseudomonas putida bearing the respective genes on plasmids. Similar work was published 10 years later by Iwakiri et al. (2004) who reported the bacterial dehalogenation of pentachloroethane to trichloroethene by an Alcaligenes sp. strain harboring an engineered P450CAM under anoxic conditions. The product was then further degraded by a genetically modified dioxygenase in the presence of oxygen. The above engineered haloaromatic dehalogenating CYP101 was expressed together with electron transport mediating elements in Sphingomonas chlorophenolicum ATCC 39723, and could deplete hexachlorobenzene (initially present at 4 mM) under transient accumulation of pentachlorophenol from a laboratory system (Yan et al., 2006). However, none of the above-mentioned genetically modified bacteria was able to grow at the expense of the xenobiotics intended to be mineralized. This means that strains transforming the target compound(s) suffer stress, require external energy sources, and may die upon short-time exposure to the compounds to be degraded. Accumulating catabolites produced during turnover of PCBs show detrimental/toxic effects (Ca´mara et al., 2004; Martı´nez et al., 2007). Within the past years solid data bases have been developed which allow the useful exploitation of the metagenome, together with the genetic tools developed for the rational construction of recombinant enzyme-encoding genes and organisms (Keasling and Bang, 1998; Meyer et al., 2007).

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Metagenomic Strategies to Mine the Enzymes of Uncultivable Bacteria

It has been estimated that more than 99% of all bacteria are uncultivable under standard laboratory conditions and therefore little accessible for research. Bacteria have been on Earth for more than 3.5 billion years and have evolved many different strategies to survive in different environments implying the existence of a huge reservoir of unknown enzyme activities including those of interest for bioremediation. It has been observed that similar dehalogenase sequences were detected in isolates from different enrichment studies but they were not the most abundant dehalogenase sequences identified in whole genome sequencing projects and massive random sequencing. On the other hand, many genes of uncertain function have been found. Thus, it appears that enrichment techniques explore a different segment of sequence space than massive sequencing of environmental DNA (Janssen et al., 2005). As a result as metagenomic technology matures an increasing number of novel enzymes are being identified. For instance, Henning et al. (2006) identified novel benzoylformate decarboxylases from a soil metagenomic library. By using stable isotope probing of soil in a pine root zone contaminated with polychlorinated biphenyls (PCBs), Leigh et al. (2007) found novel aromatic ring hydroxylating dioxygenase (ARHD) sequences which cluster distantly from all known ARHDs. On the other hand, Wexler et al. (2005) found in a waste water metagenome a novel enzyme which resembles the AdhE alcohol dehydrogenase which catalyzes the same reaction but unlike its counterparts is not inactivated by oxygen. In this way, metagenomics based technologies will furnish novel enzymes which are more difficult to find via standard enrichment studies and cultivation. However, many obstacles remain for finding such enzymes in metagenomes such as poor expression in non homologous hosts, inadequate screening systems to find interesting catabolic activities especially if they require multi-enzymatic pathways. Nevertheless, as better strategies are being developed no doubt metagenomics will provide a rich source for catabolic enzymes for degrading pollutants.

4

Improved Catabolic Strains

In order to create bacterial strains with new/improved catabolic capabilities for the efficient breakdown of environmental pollutants, improved catabolic strains are required to host catabolic pathways of interest. The starting point for the design of such catabolic strains was described in a patent filed by A. Chakrabarty of a P. putida strain harboring four catabolic plasmids for improved PAH degradation (Kellogg et al., 1981). Conjugation techniques were then successfully applied for the interspecies transfer of catabolic plasmids and other transferable genetic elements such as transposons (Reineke, 1998). Other authors also suggested strategies for the construction of improved bacterial strains with better PCB degradation capacities (Brenner et al., 1994). Also attempts have been made to improve strains to degrade other pollutants (de Lorenzo, 1994), such as 2-chlorotoluene. While the other two monochlorotoluene isomers as well as several isomers of dichlorotoluene can be mineralized and serve as carbon and energy sources for bacteria isolated from contaminated environments, 2-chlorotoluene remains recalcitrant. Although this compound can be cooxidized, even genetically engineered strains failed to grow on it due to the accumulation of misrouted intermediates (Haro and de Lorenzo, 2001; Pollmann et al., 2005).

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Therefore a caveat for the rational design of bacterial strains with new/improved catabolic capabilities for the efficient breakdown of environmental pollutants is the difficulty of obtaining new bacterial hybrids capable of efficiently degrading novel (toxic, etc.) compounds of environmental impact as new carbon- and energy sources. Simple transfer of catabolic genes into new hosts does not necessarily furnish hybrids able to grow with them, even if all theoretically required elements be present. Probably, new genes have to accommodate themselves over a longer period of time and they as well as the new host strain have to undergo an additional adaptation process until efficient expression in the desired way. In general, transfer rates are quite low and, depending on the potential new hosts, the transferred foreign DNA may undergo restriction (Murray, 2002) before becoming productively expressed. New gene ferries evading such types of immigration control should be promising tools for future genetic improvements (Tock and Dryden, 2005). One way to overcome such problems is to stably introduce novel catabolic capabilities into the genome of bacterial strains already capable of degrading related compounds. In this way a Sphingomonas sp. already able to grow with carbofuran received a methyl parathion hydrolase gene allowing the resulting strain to grow with both carbofuran and methyl parathion simultaneously (Jiang et al., 2007). Similarly, Wittich and Wolff (2007) reported the improvement of the biphenyl degrading Cupriavidus necator H850 (former Alcaligenes eutrophus -> Ralstonia eutropha -> Wautersia eutropha) to become capable of growing with commercial PCB mixtures such as Aroclor 1221 and 1232 as the only carbon and energy sources. To achieve this, the parent strain was equipped with a cassette based on a minitransposon gene ferry (de Lorenzo and Timmis, 1994), containing a regulated halocatechol pathway comprising four catabolic enzymes two of which catalyze dehalogenations (Klemba et al., 2000). Further transfer of a 2-halobenzoate dioxygenase, and an alkylbenzoate dioxygenase, the latter responsible for the dioxygenation of intermediates with bulky side groups and highly halogenated benzoates, extended the substrate range to allow the modified organism to productively degrade PCBs as depicted in > Fig. 1.

5

Microbial Releases

In order for any engineered strain to have any applications to clean-up contaminated sites, they need to be released into the environment (de Lorenzo, 1994). Here two major problems arise. Firstly, GMOs rarely survive the harsh conditions they find in the field or they fail to compete with better adapted indigenous strains for nutrients. For the same reason, more often than not the release of GMOs into the environment fail to bioremediate contaminated sites. Therefore, improved catabolic strains will also have to show fitness in the environment. Secondly, the fate of released genetically modified microorganisms and more specifically of the engineered genes remains largely unknown. As a result genetically engineered strains are required to possess highly specific and fully individual markers (Prosser, 1994) to allow detection and tracking of GMOs once liberated into the environment. Increased public concern has been drawn towards drug resistant pathogens often termed ‘‘superbugs’’ by the press. Fear that such pathogens could be somehow be related to GMO release by genetic recombination with indigenous microorganisms has led to much investigation to mitigate the risk of horizontal gene transfer thereby decreasing the risk of unwanted effects such as the dispersal of recombinant DNA (see Davison, 2005 for a review), as well as to the development of several strategies for bacterial gene containment and suicide systems (Duque et al., 1992; Molin et al., 1993; Molina et al., 1998).

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. Figure 1 On the right, the genetic elements introduced into parent strain Cupriavidus necator H-850 are shown, which confer to the generated hybrid, C. necator RW112, the new capability for growth on industrial mixtures of PCBs such as Aroclors 1221 and 1232.

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Numerous investigations in the past decades have shown that, on the one hand, catabolic elements encoded on plasmids, transposons, etc., are subject of normal gene transfer taking place in most of the environmental niches (Bogdanova et al., 1998; Ravatn et al., 1998). On the other hand there is no proof for detrimental environmental or even toxic effects upon dispersal of such genes in the examined (artificial) environments. This may provoke the stringent request for better catabolically improved biodegraders and their release in the absence of control systems so that they get distributed and further adapted, undergo further mutational modifications and random selection. The final biological products of such a process may help us in decontaminating polluted areas but also contribute to self-healing natural processes.

6

Research Needs

Apart from new catabolic genes with interesting novel features to be discovered by gene mining within metagenomic approaches, an improved understanding of evolutionary processes is required, especially with regard to the efficient and productive integration of new catabolons or single catabolic genes into the chromosome of the bacterial host cells.

References Bogdanova ES, Bass IA, Minakhin LS, Petrova MA, Mindlin SZ, Volodin AA, Kalyaeva ES, Tiedje JM, Hobman JL, Brown NL, Nikiforov VG (1998) Horizontal spread of mer operons among grampositive bacteria in natural environments. Microbiology 144: 609–620. Brenner V, Arensdorf JJ, Focht DD (1994) Genetic construction of PCB degraders. Biodegradation 5: 359–377. Ca´mara B, Herrera C, Gonza´lez M, Couve E, Hofer B, Seeger M (2004) From PCBs to highly toxic metabolites by the biphenyl pathway. Environ Microbiol 6: 842–850. Cases I, de Lorenzo V (2005) Genetically modified organisms for the environment: stories of success and failure and what we have learned from them. Int Microbiol 8: 213–222. Chen X, Christopher A, Jones JP, Bell SG, Guo Q, Xu F, Rao Z, Wong L-L (2002) Crystal structure of the F87W/Y96F/V247L mutant of cytochrome P450cam with 1,3,5-trichlorobenzene bound and further protein engineering for the oxidation of pentachlorobenzene and hexachlorobenzene. J Biol Chem 277: 37519–37526. Clement P, Matus V, Cardenas L, Gonzalez B (1995) Degradation of trichlorophenols by Alcaligenes eutrophus JMP134. FEMS Microbiol Lett 127: 51–55. Davison J (2005) Risk mitigation of genetically modified bacteria and plants designed for bioremediation. J Ind Microbiol Biotechnol 32: 639–650.

de Lorenzo V (1994) Designing microbial systems for gene expression in the field. Trends Biotechnol 12: 365–371. de Lorenzo V, Timmis KN (1994) Analysis and construction of stable phenotypes in gram-negative bacteria with Tn5- and Tn10-derived minitransposons. Methods Enzymol 235: 368–405. Duque E, Ramos-Gonzalez M, Delgado A, Contreras A, Molin S, Ramos JL (1992) Genetically engineered Pseudomonas strains for mineralization of aromatics: survival, performances, gene transfer, and biological containment. In Controlled Delivery of Crop-Protection Agents. RM Wilkins (ed.). London: Taylor & Francis, pp. 429–437. Furukawa K (2003) ‘Super bugs’ for bioremediation. Trends Biotechnol 21: 187–190. Furukawa K, Suenaga H, Goto M (2004) Biphenyl dioxygenases: functional versatilities and directed evolution. J Bacteriol 186: 5189–5196. Haro M-A, de Lorenzo V (2001) Metabolic engineering of bacteria for environmental applications: construction of Pseudomonas strains for biodegradation of 2-chlorotoluene. J Biotechnol 85: 103–113. Henning H, Leggewie C, Pohl M, Mu¨ller M, Eggert T, Jaeger K-E (2006) Identification of novel benzoylformate decarboxylases by growth selection. Appl Environ Microbiol 72: 7510–7517. Hur H-G, Sadowsky MJ, Wackett LP (1994) Metabolism of chlorofluorocarbons and polybrominated compounds by Pseudomonas putida G786(pHG-2) via

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an engineered metabolic pathway. Appl Environ Microbiol 60: 4148–4154. Iwakiri R, Yoshihira K, Ngadiman, Futagami T, Goto M, Furukawa K (2004) Total degradation of pentachloroethne by an engineered Alcaligenes strain expressing a modified camphor mono-oexygenase and a hybrid dioxygenase. Biosci Biotechnol Biochem 68: 1353–1356. Janssen DB, Dinkla IJT, Poelarends GJ, Terpstra P (2005) Bacterial degradation of xenobiotic compounds: evolution and distribution of novel enzyme activities. Environ Microbiol 7: 1868–1882. Jiang J, Zhang R, Li R, Gu J-D, Li S (2007) Simultaneous biodegradation of methyl parathion and carbofuran by a genetically engineered microorganism constructed by mini-Tn5 transposon. Biodegradation 18: 403–412. Keasling JD, Bang SW (1998) Recombinant DNA techniques for bioremediation and environ-mentallyfriendly synthesis. Curr Opin Biotechnol 9: 135–140. Kellogg ST, Chatterjee DK, Chakrabarty AM (1981) Plasmid-assisted molecular breeding: new technique for enhanced biodegradation of persistent toxic chemicals. Science 214: 1133–1135. Kimbara K (2005) Recent developments in the study of microbial aerobic degradation of polychlorinated biphenyls. Microb Environ 20: 127–134. Klemba M, Jakobs B, Wittich R-M, Pieper DH (2000) Chromosomal integration of tcb chlorocatechol pathway genes as a means of expanding the growth substrate range of bacteria to include haloaromatics. Appl Environ Microbiol 66: 3255–3261. Leigh MB, Pellizari VH, Uhlı´k O, Sutka R, Rodrigues J, Ostrom NE, Zhou J, Tiedje JM (2007) Biphenylutilizing bacteria and their functional genes in a pine root zone contaminated with polychlorinated biphenyls (PCBs). ISME J 1: 134–148. Martı´nez P, Agullo´ L, Herna´ndez M, Seeger M (2007) Chlorobenzoate inhibits growth and induces stress proteins in the PCB-degrading bacterium Burkholderia xenovorans LB400. Arch Microbiol 188: 289–297. Meyer A, Pellaux R, Panke S (2007) Bioengineering novel in vitro metabolic pathways using synthetic biology. Curr Opin Microbiol 10: 246–253. Molin S, Boe L, Jensen LB, Kristensen CS, Givskov M, Ramos JL, Bej AK (1993) Suicidal genetic elements and their use in biological containment of bacteria. Annu Rev Microbiol 47: 139–166.

Molina L, Ramos C, Ronchel MC, Molin S, Ramos JL (1998) Construction of an efficient biologically contained Pseudomonas putida strain and its survival in outdoor assays. Appl Environ Microbiol 64: 2072–2078. Murray NE (2002) Immigration control of DNA in bacteria: self versus non-self. Microbiology 148: 3–20. Pieper DH (2005) Aerobic degradation of polychlorinated biphenyls. Appl Microbiol Biotechnol 67: 170–191. Pollmann K, Wray V, Pieper DH (2005) Chloromethylmuconolactones as critical metabolites in the degradation of chloromethylcatechols: recalcitrance of 2-chlorotoluene. J Bacteriol 187: 2332–2440. Prosser JL (1994) Molecular marker systems for detection of genetically engineered microorganisms in the environment. Microbiology 140: 5–17. Ravatn R, Zehnder AJB, van der Meer JR (1998) Low frequency horizontal transfer of an element containing the chlorocatechol degradation genes from Pseudomonas sp. strain B13 to Pseudomonas putida F1 and to indigenous bacteria in laboratory-scale activated-sludge microcosms. Appl Environ Microbiol 64: 2126–2132. Reineke W (1998) Development of hybrid strains for the mineralization of chloroaromatics by patchwork assembly. Annu Rev Microbiol 52: 287–331. Tock MR, Dryden DTF (2005) The biology of restriction and anti-restriction. Curr Opin Microbiol 8: 466–472. Wackett LP, Sadowsky MJ, Newman LN, Hur H-G, Li S (1994) Metabolism of polyhalogenated compounds by a genetically engineered bacterium. Nature 368: 627–629. Wexler M, Bond PL, Richardson DJ, Johnston AWB (2005) A wide host-range metagenomic library from a wastewater treatment plant yields a novel alcohol/aldehyde dehydrogenase. Environ Microbiol 7: 1917–1926. Wittich R-M, Wolff P (2007) Growth of the genetically engineered strain Cupriavidus necator RW112 with chlorobenzoates and technical chlorobiphenyls. Microbiology 153: 186–195. Yan D-Z, Liu H, Zhou N-Y (2006) Conversion of Sphingobium chlorophenolicum ATCC 39723 to a hexachlorobenzene degrader by metabolic engineering. Appl Environ Microbiol 72: 2283–2286.

Part 6

Functional Genomics (the Paradigms) Section Editor: Victor de Lorenzo

32 Bioinformatic, Molecular and Genetic Tools for Exploring Genome-wide Responses to Hydrocarbons O. N. Reva1 . B. Tu¨mmler 2 Biochemistry Department, Bioinformatics and Computational Biology Unit, University of Pretoria, Hillcrest, Pretoria, South Africa [email protected] 2 Klinische Forschergruppe OE 6711, Medizinische Hochschule Hannover, Hannover, Germany [email protected]

1

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1258 1.1 In Silico Analyses of Genes and Oligonucleotide Signatures . . . . . . . . . . . . . . . . . . . . . . . 1258 1.2 Omics Analyses of Cellular Constituents in Man-Made Standardized Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1259 2

Genetic and Metagenomic Tools for the Analysis of the Response of Microbial Communities to Hydrocarbons in Artificial and Natural Habitats . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1260

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Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1262

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Abstract: The response profiles of bacteria to hydrocarbons in the wild can be directly assessed by high-throughput cDNA sequencing of metagenomes, tracking the fate or metabolism of labeled cells in the microbial community or can be indirectly inferred from the screening of mutant libraries for key genetic determinants. Transcriptome, proteome and metabolome data are collected from homogeneous bacterial populations that are exposed to hydrocarbons under strictly controlled culturing conditions.

1

Introduction

The analysis of the genome-wide responses of a microorganism to hydrocarbons can be divided into two tasks. First, one should address the issue whether the microorganism of interest is capable of metabolizing alkanes. This task is accomplished by data mining of genomic sequences (if available), homology-driven cloning and sequencing and straightforward in vitro tests of substrate degradation. Second, any microorganism can be investigated in its global response to hydrocarbons by applying the current omics technologies. One should note, however, that the global profiling of mRNA transcripts, proteins and metabolites will only yield meaningful data if the bacteria are exposed to hydrocarbons in meticulously controlled environments such as chemostats. If one wants to study the responses of microbial communities to hydrocarbons in the wild, genetic or metagenomic approaches should be pursued that are adapted to the particular habitat on a case-to-case basis.

1.1

In Silico Analyses of Genes and Oligonucleotide Signatures

Genes encoding enzymes for alkane degradation can be detected by the criteria of protein sequence similarity, operon organization and conserved protein domains. An established tool is the BlastP algorithm (http://www.ncbi.nlm.nih.gov/blast/). Knowledge about the abundance of these genes is catalogued in public databases such as BRENDA (http://www. brenda-enzymes.info/) (Barthelmes et al., 2007). A more demanding task is the search for the promoters. Promoter sequences of alkane degradation operons show rather weak homology that hinders their prima facie identification. The promoter sequences may however be indirectly detected by analysis of oligonucleotide composition. Promoter regions typically exhibit higher DNA curvature, lower base stacking energy and are more rigid (Reva et al., 2008). The z-score structural profiles for the promoter regions (usually starting from 400 bp upstream to gene start codon) may be compared with the standard profiles published at www.cbs.dtu.dk/services/GenomeAtlas/. Consensus motifs of promoter regions may be identified by the MEME program (http://meme.nbcr.net/meme/ intro.html) (Bailey et al., 2006). Horizontal gene transfer contributed to the spread of alkane metabolic activities among g-Proteobacteria. Besides the search for mobile genetic elements within or adjacent the alkane degradation operons (van Beilen et al., 2001), evidence for horizontal gene transfer is gained from the comparison of oligonucleotide compositional biases between the gene cluster and the surrounding chromosomal sequences (> Fig. 1). In the A. borkumensis SK2 genome, for example, the two regions comprising alkB1 and alkB2 genes are as similar to each other in their tetranucleotide usage (TU) patterns as each of them is to Yersinia species. The two alk

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. Figure 1 Alternative oligonucleotide usage in the gene cluster encoding proteins of the alkane degradation pathway in the Alcanivorax borkumensis SK2 genome. The genomic fragment [3,060,396–3,068,240] of A. borkumensis SK2 comprising alkB1 regulator, alkB1, alkH1, alkH2 and alkJ is highlighted in yellow on the 80 kb window of the genomic gene map. The alkane degradation gene cluster is characterized by below-average GC-content (black line, upper panel) and an atypical tetranucleotide usage (TU). The low blue line shows the variations of local TU patterns from the genomic average. The dotted lines indicate a 50% G + C-content (upper panel) and the average deviation of local TU (8 kb window) from the genomic average (lower panel). The oligonucleotide usage compositional biases were analyzed by the SeqWord Genome Browser (Ganesan et al., 2008) which is publicly accessible online at http://www.bi.up.ac.za/ SeqWord/or http://genomics1.mh-hannover.de/seqword/.

genes, however, differ in their TU usage from the bulk sequence of A. borkumensis indicating that the alkB1 and alkB2 genes were delivered to A. borkumensis from an ancestor of the Yersinia lineage (Reva et al., 2008).

1.2

Omics Analyses of Cellular Constituents in Man-Made Standardized Environments

Transcriptome, proteome and metabolome analyses can generate comprehensive quantitative profiles of the genome-wide response of a microorganism to hydrocarbons. All these omics technologies determine average values of bulk samples containing millions of bacteria. Consequently the most meaningful data are produced from homogeneous bacterial populations with minimal variation of the expression profiles of individual cells. To minimize spatiotemporal gradients, bacteria should be grown under carefully controlled conditions in batch cultures or – better though – in chemostats. In the latter case stationary cultures are perturbed by the stimulus of interest, i.e., in this context the exposure to hydrocarbons. Experiments in chemostats will yield reproducible and reliable quantitative data with low statistical spread as it is requested for applications in systems biology or white biotechnology.

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Global mRNA expression profiles are typically obtained by the hybridization of bacterial cDNA onto Gene chips that have each open reading frame of the genome represented by either one long (50–70 mers) or numerous pairs of short complementary and single mismatch oligonucleotides (20–35 mers) (Dharmadi and Gonzalez, 2004; Ness, 2007). The experimental protocols comprise the RNA preparation from bacterial cells, cDNA synthesis, fragmentation, and labeling and the hybridization of the cDNA preparation onto the microarray. After washing off non-specific binding, the hybridization signals are scanned and processed. A time series experiment of the influence of hydrocarbons on the bacterial mRNA expression profile will typically require 3–4 biological replicates per time point. Particular care should be taken for the proper normalization of signals of gene probesets and the non-trivial identification of up- and downregulated genes. An acceptable rate of false positive signals is calculated by permutation methods like The Significance Analysis of Microarrays (SAM) (freely available at http://www-stat.stanford.edu/tibs/SAM/). In addition to global mRNA expression profiling, the ‘‘ChIP-chip’’ approach (Liu, 2007) may be applied to map the binding and regulatory genomic sites of transcription factors that regulate the bacterial response to hydrocarbons. The transcription factor is crosslinked with genomic DNA, fragmented to approximately 500 bp and immunoprecipitated with an antibody. The coprecipitated DNA is hybridized on a genome-spanning tiling microarray and the genomic map positions of the transcription factor binding sites are identified from the hybridization signals of the positive probes. Considering the short average lifetime of a bacterial mRNA of less than 2 min, the transcriptome will provide a snapshot of the global expression profile at that particular point of time. The longer living proteins visualize the cellular response to a signal such as hydrocarbons in a broader time-frame. With the emergence of mass spectrometry (MS) in protein science and the availability of complete genome sequences, bacterial proteomics has gone through a rapid development. The application of gel-based and gel-free technologies, the analyses of subcellular proteome fractions (Cordwell, 2006) and the use of multidimensional capillary HPLC combined with MS/MS have allowed high qualitative and quantitative coverage of currently more than one-third of the theoretical bacterial proteome (Wolff et al., 2007; Xia et al., 2007). Metabolomics is the youngest omics discipline that analyzes metabolic profiles in response to environmental compounds and signals. Metabolites are a chemically highly diverse group of compounds. Hence the analysis of microbial metabolomes is a formidable challenge. Protocols need to be more flexible than those for proteomics or transcriptomics. Gas chromatography, liquid chromatography or capillary electrophoresis are combined with numerous MS methods (Dunn, 2008). State of the art metabolomics platforms detect up to 1,000 different metabolites but only a minority of which may be identifiable because the mass spectra of metabolites and their volatile derivatives often are not known or not listed in the available spectral libraries. In summary, metabolomics is a young and vibrant scientific discipline with workflows are just being developed leaving still much room for improvement.

2

Genetic and Metagenomic Tools for the Analysis of the Response of Microbial Communities to Hydrocarbons in Artificial and Natural Habitats

The investigation of heterogeneous microbial communities requires the spatiotemporal resolution of the signals of individual cells. To visualize the growth and decay of individual species

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in the population upon exposure to hydrocarbons, the target organisms may be labeled with specific fluorescence markers, for example ribosomal ribonucleic acid targeted oligonucleotide probes (Daims and Wagner, 2007). Labeled cells are visualized by fluorescence microscopy and are quantified by direct visual cell counting or by digital image analysis. To get an estimate of the global growth activity in the absence and presence of hydrocarbons, recombinant marker cells may be engineered that harbor a fusion of a ribosomal rrn promoter to a gene encoding an unstable variant of the green fluorescent protein (GFP) or equivalent (Ramos et al., 2000). Ribosomal contents and synthesis rates are thus monitored by the fluorescence of GFP-positive cells. As the next step, one may study the metabolic activity of the community in more depth. Stable isotopes or radioisotopes can be incorporated into bacterial sub-populations (Neufeld et al., 2007). Subsequent analysis of labeled biomarkers of sub-populations with stable-isotope probing (DNA, RNA or phospholipid-derived fatty acid) or of individual cells with a combination of fluorescence in situ hybridization and microautoradiography reveals linked phylogenetic and functional information about the organisms that assimilated the compounds of interest such as hydrocarbons. A complementary approach to the analysis of metabolic activities is the genome-wide search for the key determinants of the bacterial response to hydrocarbons by In Vivo Expression Technology (IVET) (Rediers et al., 2005), Signature-Tagged Mutagenesis (STM) (Mazurkiewicz et al., 2006), differential display using arbitrarily primed PCR (Fislage, 1998), subtractive and differential hybridization (Ito and Sakaki, 1997) or selective capture of transcribed sequences (SCOTS) (Graham and Clark-Curtiss, 1999). We discuss the most widely used IVET and STM strategies. IVET involves the construction of a conditionally compromised strain that is mutated in a gene encoding an essential growth factor (egf). This mutant strain is not able to grow in the environment under study. The second component of IVET is the promoter trap, consisting of a promoterless egf gene and a transcriptionally linked reporter gene (rep). Bacterial DNA is cloned randomly into the promoter trap and integrated in the chromosome of the egf mutant strain. Only in strains that carry a promoter active in the specified niche can the egf mutation be complemented. After selection in this environment, bacteria are reisolated and spread on a general growth medium that is suitable for monitoring reporter gene activity in vitro. Accordingly, constitutive promoters are distinguished from promoters that are specifically induced in the wild. Colonies bearing the latter type of transcriptional fusion are subjected to a second IVET screening to eliminate false positives. STM is a mutation-based screening method that uses a population of isogenic transposon mutants for the identification of essential genes by negative selection. A pool of mutants can simultaneously be examined because they are differentiated by unique DNA marker sequences, the ‘‘signature tags.’’ The pools of mutants are exposed to the habitat of interest. One screens for mutants that are unable to survive or to grow because they are inactivated in a key gene for survival in this habitat of interest. The beauty of the technology is an in vivo selection process done by the habitat among a mixed population of mutants. The studies of genome-wide responses in bacteria will soon experience a quantum leap by next-generation sequencing technologies (Schuster, 2008). The expression profile of a bacterial metagenome will be determined from straightforward ultra-high throughput cDNA sequencing (Huson et al., 2007; Urich et al., 2008) and simultaneously the presence and abundance of species in the metagenome are estimated from the occurrence of informative marker oligonucleotides in the primary sequence data (Davenport et al., 2008).

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Research Needs

Next-generation sequencing technologies will dominate genomics within the foreseeable future. Sequencing capacities may soon outdistance computing capacities. Soft- and hardware will have to be developed for storage and handling of the anticipated avalanche of primary data. New and sophisticated algorithms will be needed for data mining and interpretation. Within the omics and systems biology fields, comprehensive static and dynamic descriptions of cellular constituents will only become feasible with more sophisticated methodologies. Transcriptomics has reached a mature state, but proteomics and particularly metabolomics still require substantial improvements in detection and identification rates.

References Bailey TL, Williams N, Misleh C, Li WW (2006) MEME: discovering and analyzing DNA and protein sequence motifs. Nucleic Acids Res 34: W369–W373. Barthelmes J, Ebeling C, Chang A, Schomburg I, Schomburg D (2007) BRENDA, AMENDA and FRENDA: the enzyme information system in 2007. Nucleic Acids Res 35 (Database issue): D511–D514. Cordwell SJ (2006) Technologies for bacterial surface proteomics. Curr Opin Microbiol 9: 320–329. Daims H, Wagner M (2007) Quantification of uncultured microorganisms by fluorescence microscopy and digital image analysis. Appl Microbiol Biotechnol 75: 237–248. Davenport CF, Wiehlmann L, Reva ON, Tu¨mmler B (2009) Analysis of Pseudomonas genomes by abundant 8–14-mer oligonucleotides. Environ Microbiol 11. DOI:10.1111/j.1462-2920.2008.01839.x. Dharmadi Y, Gonzalez R (2004) DNA microarrays: experimental issues, data analysis, and application to bacterial systems. Biotechnol Prog 20: 1309–1324. Dunn WB (2008) Current trends and future requirements for the mass spectrometric investigation of microbial, mammalian and plant metabolomes. Phys Bio 5: 11001. Fislage R (1998) Differential display approach to quantitation of environmental stimuli on bacterial gene expression. Electrophoresis 19: 613–616. Ganesan H, Rakitianskaia AS, Davenport CF, Tu¨mmler B, Reva ON (2008) The SeqWord Genome Browser: an online tool for the identification and visualization of atypical regions of bacterial genomes through oligonucleotide usage. BMC Bioinformatics 9: 333. Graham JE, Clark-Curtiss JE (1999) Identification of Mycobacterium tuberculosis RNAs synthesized in response to phagocytosis by human macrophages by selective capture of transcribed sequences (SCOTS). Proc Natl Acad Sci USA 96: 11554–11559.

Huson DH, Auch AF, Qi J, Schuster SC (2007) MEGAN analysis of metagenomic data. Genome Res 17: 377–386. Ito T, Sakaki Y (1997) Fluorescent differential display. Methods Mol Biol 85: 37–44. Liu XS (2007) Getting started in tiling microarray analysis. PLoS Comput Biol 3: 1842–1844. Mazurkiewicz P, Tang CM, Boone C, Holden DW (2006) Signature-tagged mutagenesis: barcoding mutants for genome-wide screens. Nat Rev Genet 7: 929–939. Ness SA (2007) Microarray analysis: basic strategies for successful experiments. Mol Biotechnol 36: 205–219. Neufeld JD, Wagner M, Murrell JC (2007) Who eats what, where and when? Isotope-labelling experiments are coming of age. ISME J 1: 103–110. Ramos C, Mølbak L, Molin S (2000) Bacterial activity in the rhizosphere analyzed at the single-cell level by monitoring ribosome contents and synthesis rates. Appl Environ Microbiol 66: 801–809. Rediers H, Rainey PB, Vanderleyden J, De Mot R (2005) Unraveling the secret lives of bacteria: use of in vivo expression technology and differential fluorescence induction promoter traps as tools for exploring niche-specific gene expression. Microbiol Mol Biol Rev 69: 217–261. Reva ON, Hallin PF, Willenbrock H, Sicheritz-Ponten T, Tu¨mmler B, Ussery DW (2008) Global features of the Alcanivorax borkumensis SK2 genome. Environ Microbiol 10: 614–625. Schuster SC (2008) Next-generation sequencing transforms today’s biology. Nat Methods 5: 16–18. Urich T, Lanze´n A, Qi J, Huson DH, Schleper C, Schuster SC (2008) Simultaneous assessment of soil microbial community structure and function through analysis of the meta-transcriptome. PLoS ONE 3: e2527. van Beilen JB, Panke S, Lucchini S, Franchini AG, Ro¨thlisberger M, Witholt B (2001) Analysis of Pseudomonas putida alkane-degradation gene clusters

Bioinformatic, Molecular and Genetic Tools and flanking insertion sequences: evolution and regulation of the alk genes. Microbiology 147: 1621–1630. Wolff S, Antelmann H, Albrecht D, Becher D, Bernhardt J, Bron S, Bu¨ttner K, van Dijl JM, Eymann C, Otto A, Tam le T, Hecker M (2007) Towards the entire proteome of the model bacterium Bacillus

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subtilis by gel-based and gel-free approaches. J Chromatogr B Analyt Technol Biomed Life Sci 849: 129–140. Xia Q, Hendrickson EL, Wang T, Lamont RJ, Leigh JA, Hackett M (2007) Protein abundance ratios for global studies of prokaryotes. Proteomics 7: 2904–2919.

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33 Alcanivorax borkumensis V. Martins dos Santos1 . J. Sabirova2 . K. N. Timmis2 . M. M. Yakimov3 . P. N. Golyshin2,4 1 Systems and Synthetic Biology Research Group, Helmholtz Center for Infection Research, Braunschweig, Germany [email protected] 2 Environmental Microbiology Laboratory, Helmholtz Centre for Infection Research, Braunschweig, Germany 3 School of Biological Sciences, Bangor University, Gwynedd, UK 4 Istituto per l’Ambiente Marino Costiero, CNR, Messina, Italy 1 1.1 1.2 1.3

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1266 Alcanivorax: A Cosmopolitan Paradigm of Hydrocarbonoclastic Bacteria . . . . . . . . . . 1266 Phylogenetic Placement of Alcanivorax . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1266 Choice of Alcanivorax for Functional Genomic and Physiologic Characterization . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1267

2 2.1 2.2 2.3

Genome Sequence of Alcanivorax borkumensis SK2 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1267 Global Features of the A. borkumensis SK2 Genome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1267 Genomic Islands and Mobile Genetic Elements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1268 Genomic Comparisons of A. borkumensis SK2 to Related Species . . . . . . . . . . . . . . . . . . 1269

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Genomic Basis of Alkane Degradation in A. borkumensis SK2 . . . . . . . . . . . . . . . . . 1271

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The Genetic Potential of A. borkumensis SK2 for the Emulsification of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1272

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Biofilm Formation, Cell Surface, and Secretion in A. borkumensis SK2 . . . . . . . . 1275

6 6.1 6.2 6.3

Nutrient Transport, Regulation, and Stress Responses . . . . . . . . . . . . . . . . . . . . . . . . . . . 1277 Nutrient Transport and Sodium Dependency . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1277 Stress Responses and Osmoregulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1278 Regulation and Signal Transduction in A. borkumensis SK2 . . . . . . . . . . . . . . . . . . . . . . . . 1280

7 Metabolic Specialization in Oligotrophic Environments . . . . . . . . . . . . . . . . . . . . . . . . . 1281 7.1 Glyoxylate Bypass and Gluconeogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1282 8 Genome-Scale, Constraint-Based Modeling of the A. borkumensis Metabolism 1283 8.1 Storage Compounds and Polyhydroxyalkanoate Production by A. borkumensis . . . 1284 9

Concluding Remarks and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1285

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Alcanivorax borkumensis

Abstract: Alcanivorax borkumensis is a marine bacterium that uses exclusively petroleum oil hydrocarbons as sources of carbon and energy (and is therefore designated ‘‘hydrocarbonoclastic’’). It is found in low numbers in all oceans of the world and becomes abundant in oil-contaminated waters. Its ubiquity, unusual physiology and demonstrated role in biodegradation show that it is globally important in the removal of hydrocarbons from polluted marine systems. Genome sequencing, extensive functional genomic analysis and genome-wide constraint-based modeling of the metabolism Alcanivorax borkumensis SK2 type strain, an outstanding paradigm of hydrocarbonoclastic bacteria, has provided substantial insights into the genomic basis of the efficiency and versatility of its hydrocarbon utilization, nutrient scavenging capabilities, niche-specific stress responses and the metabolic routes and flux distributions underlying its remarkable hydrocarbon utilization abilities. The wealth of information thus far generated provides a solid knowledge-base for the understanding the physiology and ecological success of this fascinating and globally important bacterium and for the design of new strategies to mitigate the ecological damage caused by oil spills.

1

Introduction

1.1

Alcanivorax: A Cosmopolitan Paradigm of Hydrocarbonoclastic Bacteria

In 1998 Yakimov et al. (1998) reported the isolation from a North Sea sediment, and characterization, of Alcanivorax borkumensis, an unusual marine bacterium able to grow on a highly restricted spectrum of substrates, predominantly alkanes. Within a short period of time, the isolation of Alcanivorax, or detection of its 16S rRNA gene sequences, from samples taken from the Atlantic Ocean, Mediterranean Sea, North Sea, Sea of Japan, South China Sea, and the Antarctic was reported, so it is a truly cosmopolitan bacterium. An updated biogeography of Alcanivorax borkumensis strains and other marine obligately hydrocarbonoclastic bacteria (OHCB) is described in more detail by Cappello & Yakimov (See > Chapter 6, Vol. 3, Part 1), respectively. Significantly, Alcanivorax is found in low numbers in unpolluted waters, but in high abundances in oil-polluted waters and coastlines, where it may comprise 80–90% of the oil-degrading microbial community (Harayama et al., 1999; Kasai et al., 2001). Moreover, Alcanivorax becomes abundant in field and mesocosm experiments involving the addition of nitrogen and phosphorus fertilizers to stimulate microbial degradation of oil (Kasai et al., 2002; Ro¨ling et al., 2002, 2004, Cappello et al., 2007, Capello & Yakimov, volume 4, > Chapter 11, Vol. 5, Part 1). These findings indicate that Alcanivorax plays a critical role in the natural cleansing of oil-polluted marine systems, and implies that it could constitute the basis of novel biotechnology strategies to accelerate the environmental repair process.

1.2

Phylogenetic Placement of Alcanivorax

A. borkumensis is classified within the order Oceanospirillales of the g-subclass of the Proteobacteria. The type strain of genus Alcanivorax is North Sea isolate A. borkumensis SK2 (DSM 11573T). Since its isolation, 15 other species have thus far been described, including

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Alcanivorax jadensis DSM 12178T and Alcanivorax venustensis DSM 13974T (Bruns and Berthe-Corti, 1999; Ferna´ndez-Martı´nez et al., 2003), Alcanivorax dieselolei DSM 16502T (Liu and Shao, 2005), and Alcanivorax balearicus MACL04T (Rivas et al., 2007). These bacteria are Gram-negative, aerobic, rod-shaped and use a limited number of organic compounds, including aliphatic hydrocarbons, volatile fatty acids, and pyruvate and its methyl ether. The 16S rRNA gene sequence analysis showed that these strains are all members of the c-subclass of the Proteobacteria. The precise phlyogenetic placement of Alcanivorax strains is described in > Chapter 6, Vol. 3, Part 1.

1.3

Choice of Alcanivorax for Functional Genomic and Physiologic Characterization

Its global importance, its unusual physiology, and its potential for biotechnological applications, made Alcanivorax borkumensis the obvious choice for functional genomic analysis of a paradigm of OHCBs. The aim was to gain insights into the genomic basis of its unusual metabolic capability and cellular composition, its peculiar lifestyle and high affinity for hydrocarbon substrates, its genomic responses to the signals and environmental stresses it faces in its hostile environment, its ability to degrade a range of oil hydrocarbons and to dominate oil-degrading microbial communities, and to access genomic information central to the biotechnological potential of this fascinating bacterium. The main results of the (comparative) sequence analysis of the type strain Alcanivorax borkumensis SK2 (Schneiker et al., 2006; Reva et al., 2008), its functional analysis and niche-specificity (Sabirova et al., 2006a, 2008), biotechnological potential (Sabirova et al., 2006b; Kalscheuer et al., 2007) and genome-scale metabolic modeling (Garcia et al., unpublished) are summarized below.

2

Genome Sequence of Alcanivorax borkumensis SK2

2.1

Global Features of the A. borkumensis SK2 Genome

The A. borkumensis SK2 genome consists of a single circular chromosome of 3,120,143 base pairs (bp) with an average G þ C content of 54.7%. The assembly of the sequence was validated by a complete BAC map (Schneiker et al., 2006). Biological roles were assigned to 2,241 of the 2,755 predicted coding sequences (CDS). The remaining 541 CDS comprise 316 conserved hypothetical CDS and 198 CDS of unknown function. The chromosome of Alcanivorax borkumensis SK2 is highly symmetric and homogeneous (Reva et al., 2008). The origin and terminus of replication are located opposite to each other in the chromosome and are discerned with high signal to noise ratios by maximal oligonucleotide usage biases on the leading and lagging strand. The structure of the genomic DNA is also rather uniform throughout the chromosome with respect to properties such intrinsic curvature, repeat properties (global direct and inverted repeats, which are globally low), position preference (low position preference correlate with enrichment of highly expressed genes), or base stacking energy (which measures helix rigidity and is therefore a measure of flexibility). The chromosome contains only few regions with extraordinarily low or high curvature, position preference or base stacking energy (> Fig. 1).

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. Figure 1 Genome Atlas of A. borkumensis SK2 showing different structural parameters and the distribution of global repeats, GC skew and A þ T contents. Color intensity increases with the deviation from the average. Values close to the average are shaded very light grey; values with more than three standard deviations from the average are most strongly colored. Courtesy David W. Ussery, Danish Technical University (http://www.cbs.dtu.dk/researchgroups/ compmicro.php).

2.2

Genomic Islands and Mobile Genetic Elements

The A. borkumensis SK2 genome has 18 regions with atypical GC-content and thus predicted to be of alien origin (Schneiker et al., 2006; Reva et al., 2008). This number is relatively low number, for instance, as compared to the P. putida KT2440, with which is strongly related (see below) and which has threefold more islands per megabase than A. borkumensis. Several of atypical these regions, including a large island comprising a cluster of 40 genes, specify proteins characteristic of cell-surface polysaccharide biosynthesis. This suggests that a substantial fraction of the A. borkumensis SK2 determinants for cell surface structures may have been acquired by horizontal gene transfer. Interestingly, the gene clusters that confer the assimilation of aliphatic hydrocarbons (van Beilen et al., 2004; Yakimov et al., 2007), are localized in two genome islands which Reva et al. (2008) claim that were probably acquired from an ancestor of the Yersinia lineage. In contrast, the alk genes of Pseudomonas putida still exhibit the typical Alcanivorax oligonucleotide signature, which reflects the complex evolution of this major hydrocarbonoclastic trait.

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The A. borkumensis SK2 genome harbors also only a small number of mobile genetic elements such as transposons, IS elements or their remnants. The only putative intact IS element identified in A. borkumensis SK2, carries two transposase encoding genes tnpB (ABO_2081) and tnpA (ABO_2082) and is flanked by 16 bp perfect inverted repeats (IR). Additionally, a putative transposon carrying a heavy metal cation efflux transport system was identified. The transposon is delimited by two nearly identical, albeit incomplete IS elements. The two putative IS elements each encode, besides the putative transposase (ABO_2468 and ABO_2475, respectively), a hypothetical protein and a CzcA-like protein. Along with the czcBCD gene cluster, located in the central part of the transposon, it contains the complete set of structural genes and a regulatory gene for a heavy metal resistance cation efflux transport system. Another possibly incomplete transposon, containing two transposases (ABO_1354 and ABO_1355) was detected. Gene clusters consisting of heavy metal resistance determinants like czcABC and copABC are located downstream. This might hint to the acquisition of heavy metal resistance determinants by horizontal gene transfer. This paucity of mobile elements in A. borkumensis may reflect the relatively stable seawater environment in which it thrives and a small extent of genetic interactions.

2.3

Genomic Comparisons of A. borkumensis SK2 to Related Species

A. borkumensis strain SK2 is the first member within the order Oceanospirillales of the g-subclass of the Proteobacteria whose genome was sequenced. Most of its genes have highest similarity to annotated genes of bacteria belonging to the gamma – (1897; 68.9%) and b-subclasses (302; 11.0%) of the Proteobacteria. A. borkumensis shares a similar oligo (tetra-) nucleotide usage and promoter structure with the Pseudomonadales. Orthologue comparisons to non-marine members of the Pseudomonaceae, various marine g- and a-Proteobacteria as well as to selected marine Gram-positive bacteria revealed that the number of orthologs roughly decreases with increasing phylogenetic distance. Interestingly, despite of the variation in GC-content, the patterns of tetranucleotide usage shows that A. borkumensis forms a cluster with Pseudomonas, Methylococcus, Xanthomonas, and Xylella. This cluster is clearly separated from other g-Proteobacteria (Reva et al., 2008). To further investigate the relationship of A. borkumensis SK2 to related species, orthologue analyses were performed against bacterial species which had shown the highest number of best homologues in BlastP searches against the NCBI ‘‘nr’’ database. The compared species include non-marine members of the Pseudomonaceae and various marine bacterial strains. The orthologs and paralogs of A. borkumensis genes with the highest sequence homology were found in most cases among g-Proteobacteria, with Acinetobacter and P. aeruginosa as closest relatives. Comparisons of the predicted protein sequences of A. borkumensis SK2 (cutoff value of E-30) with the three pseudomonads yielded 1,616 (58.7%), 1,478 (53.6%) and 1,329 (48.2%) orthologs in P. aeruginosa, P. putida, P. syringae, respectively. The number of orthologues to the predicted protein sequences of A. borkumensis SK2 in marine g-Proteobacterial strain amounts: 1,205 CDS (43.7%) in S. degradans, for which only a draft genome is available, 1,233 CDS (44.8%) in V. vulnificus, 1,165 CDS (42.3%) in I. loihiensis, and 1,222 CDS (44.4%) in P. profundum. Further orthologue analyses with bacteria of marine habitats belonging to the a-Proteobacteria (Caulobacter crescentus, Silicibacter pomeroyi) and to the Gram-positive bacterium Oceanobacillus iheyensis yielded less orthologs: 931 CDS (33.8%), 917 CDS (33.3%) and 684 CDS (24.8%), respectively. These results roughly indicate that the increase of phylogenetic distance

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. Figure 2 Genetic determinants involved in alkane degradation in A. borkumensis SK2. (a) Clusters of genes coding for proteins involved in alkane degradation in A. borkumensis SK2 (see text for

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between the analyzed bacteria (> Fig. 2) is accompanied by a decrease of the number of orthologues. Therefore, a relatively conserved gene order between A. borkumensis SK2 and the pseudomonads might be expected. However, synteny analyses revealed only weak linear relationships of the A. borkumensis SK2 genome sequence with the genome sequences of P. aeruginosa and P. putida. Syntenic regions between A. borkumensis SK2 and each of the four marine g-Proteobacteria in all cases were even smaller compared to those of the pseudomonads.

3

Genomic Basis of Alkane Degradation in A. borkumensis SK2

A. borkumensis SK2’s most distinctive feature is its ability to grow efficiently and almost exclusively on alkanes (Yakimov et al., 1998, 2007, Capello & Yakimov See > Chapter 6, Vol. 3, Part 1). Consistent with this, A. borkumensis SK2 specifies a number of systems for the catabolism of hydrocarbons (> Fig. 2). The SK2 alkSB1GHJ operon genes (ABO_2707ABO_2710, > Fig. 2a) have, with the exception of alkS (coding for a LuxR positive regulator), above 80% similarity at the amino acid level to the corresponding, well-characterized alkane degradation components in P. putida strains (van Beilen et al., 2004, Yakimov et al., 2007). The SK2 AlkB1 alkane hydroxylase oxidizes efficiently medium-chain alkanes (C5 to C12). Its expression is alkane-induced in the exponential growth phase, but is not expressed in the stationary phase (van Beilen et al., 2004; Sabirova et al., 2006b). Other genes related to this alkane degradation pathway, namely, alkK (ABO_2748), alkL (ABO_1922) and alkN (ABO_0106), have lower similarities to the orthologues in P. putida, whereas no homolog could be detected for the alkF gene coding for rubredoxin 1 in P. putida GPo1. However, two genes, rubA and rubB (ABO_0163 and ABO_0162) encode for a rubredoxin and a rubredoxin reductase, respectively and are likely to be involved in alkane catabolism. Rubredoxin reductase genes are not clustered with alkane hydroxylase genes in many alkane-degrading bacteria (van Beilen et al., 2004). A second alkane hydroxylase system is specified by ABO_0121, coding for a GntR transcriptional negative regulator and ABO_0122, coding for the alkane hydroxylase AlkB2, which oxidizes medium-chain alkanes in the range C8 to C16 (van Beilen et al., 2004). Although both alkB1 and alkB2 genes in A. borkumensis strains AP1 and SK2 are simultaneously induced by C10 to C16 alkanes (van Beilen et al., 2004), their underlying regulation is certainly different and possibly responds complementarily under different conditions (van Beilen et al., 2005). Importantly, both alkane hydroxylase gene systems are located close to the origin of replication of the chromosome (> Fig. 2a), which provides in A. borkumensis cells rapidly growing on alkanes a high dosage of alkane hydroxylation genes, and presumably therefore high catabolic activity. Although the growth on isoprenoids typically showed the longer lag phase, the doubling times of exponentially growing A. borkumensis SK2 cells

details). (b) Neighborhood-joining tree showing the phylogenetic affiliation of the P450 cytochromes of A. borkumensis SK2. Two identical alkane-induced cytochromes P450(b) (ABO_2288) and P450(c) (ABO_0201) cluster with the Acinetobacter EB104 cytochrome P450. Cytochrome P450(a) (ABO_2384) is affiliated distantly. The tree is rooted with the methylCoM-reductase subunit A from Methanolubus tindarius (gb|U22244). (c) Reconstruction of putative alkane degradation pathways in A. borkumensis SK2 (see text for details).

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were comparable at the expense both of hexadecane (0.115  0.03 h1) and pristane (0.106  0.016 h1) as a single source of carbon and energy. Yakimov et al. (Denaro & Yakimov, See > Chapter 21, Vol. 4, Part 3) found experimentally that A. borkumensis is able to degrade alkanes up to C32, as well as the long-chain isoprenoids phytane and pristane and alkyl-aromatic hydrocarbons. This spectrum is much broader than expected on the basis of the current knowledge of the substrate ranges of alkane hydroxylase complexes, suggesting that the SK2 genome specifies a wider range of systems involved in hydrocarbon catabolism. It codes for three cytochromes: ABO_2384 for P450(a), and two genes ABO_2288 and ABO_0201 for the paralogous P450(b) and P450(c) (> Fig. 2a). P450(c) clusters in an operon-like structure with genes coding for a ferredoxin (fdx, ABO_0200), an alcohol dehydrogenase (alkJ2, ABO_0202), a FAD-dependent oxidoreductase (ABO_0203) and an araC transcriptional regulator gene (ABO_0199) (> Fig. 2a). A proteomic profiling study by Sabirova et al. (2006a) revealed that all the enzymes coded by this gene cluster are upregulated during growth on octadecane. Quantitative real-time transcriptional analysis showed that the two paralogous genes P450(b) and P450(c) are expressed only in the presence of alkane, whereas P450(a) is expressed constitutively (Sabirova et al., unpublished). The two alkane-induced cytochromes of SK2 are phylogenetically grouped in one branch with the P450 cytochrome of Acinetobacter sp. EB104 (> Fig. 2b), that catalyses the terminal oxidation of alkanes. The ability of these two P450 cytochromes of the CYP153 family to catalyze terminalalkane hydroxylation was experimentally tested by van Beilen et al. (2006). Altogether thus, the three P450 systems likely account for an important part of the extensive hydrocarbon degradation capacity of A. borkumensis. A metabolic reconstruction of these various alkane degradation pathways is depicted in > Figs. 2c and > 4. The existence of several different alkane-oxidizing systems is consistent with the broad range of oil hydrocarbons degraded by A. borkumensis and its ecological success in oilcontaminated marine habitats. In their proteomic profiling study, Sabirova et al. (2006a) found that, among other gene products to be up-regulated on alkanes were proteins related to the further metabolism of alkanes, like fatty acid oxidation (FadB, FadB2) and polyhydroxyalkanoate production (PhaC), a number of proteins involved in transcriptional and translational regulation processes, as well as transport proteins and a number of hypothetical and conserved hypothetical proteins of unknown function. It is anticipated that a fraction of the large number of enzymes (in particular) with unknown specificity encoded by the SK2 genome may be involved in hydrocarbon catabolism. The versatility and wider spectrum for hydrocarbon utilization provides A. borkumensis SK2 a competitive advantage over other members of oil-based marine microbial communities and underscores its potential for oil clean up in marine and coastal habitats.

4

The Genetic Potential of A. borkumensis SK2 for the Emulsification of Hydrocarbons

The production of biosurfactants facilitates emulsification of alkanes, decreases droplet size, thereby increases the hydrocarbon:water interface surface area and thus hydrocarbon bioavailability, and enhances the degradation rate of these hydrophobic organic substrates. A. borkumensis forms stable emulsions of hydrocarbon in water and produces biosurfactants (> Fig. 3). Biochemical analysis of these biosurfactants revealed their structure as anionic glucolipids carrying four fatty acids of varying chain lengths (Abraham et al., 1998). These

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. Figure 3 Transmission electron micrograph of Alcanivorax borkumensis cells growing at a water – n-hexadecane interface. The cellular shape is rather irregular and most cells contain electron-translucent inclusions, food storage granules, of different size and number (Credits: Heinrich Lu¨nsdorf at the HZI).

glucolipids exist act in two ways: (1) a glycine-containining biosurfactant precursor linked to the cell surface increases the hydrophobicity of the cells and their affinity to oil droplets suspended in the water phase, and (2) mature extracellular lipid biosurfactant is released from the cell into the surrounding medium, thereby promoting the creation of micelles with water insoluble fractions, hence increasing the bioavailability of hydrocarbons (Yakimov et al., 1998, Abraham et al., 1998). The genetic organization of the glucolipid biosynthesis remains unclear in A. borkumensis SK2, but genome annotation revealed several candidate genes potentially involved in biosurfactant production. ABO_1783 and ABO_2215 encode glycosyltransferases, exhibiting significant homology with rhlB from P. aeruginosa (Ochsner et al., 1994) and glycosyltransferase protein family 9, respectively. These gene products possibly provide the sugar moiety of the glucolipids and may be involved in the glycosylation of lipid moiety, yielding glucose lipid surfactants. A. borkumensis SK2 expresses as well an OprF/OmpA protein encoded by ABO_0822, that is up-regulated on alkane (Sabirova et al., 2006a). OmpA proteins, the active constituents of the biosurfactant alasan, have been demonstrated to efficiently emulsify

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. Figure 4 Schematic overview of metabolism and transport in A. borkumensis SK2. The background is a transmission electron micrograph (TEM) of an Alcanivorax

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hydrocarbons and to be produced by a number of oil-degrading g-Proteobacteria (Ron and Rosenberg, 2001). A. borkumensis SK2 harbors determinants of OprG/OmpW (ABO_1922) and OmpH (ABO_1152), which are also possibly involved in emulsifier production. Thus, hydrocarbons are emulsified b A. borkumensis both through the creation of stable emulsions

5

Biofilm Formation, Cell Surface, and Secretion in A. borkumensis SK2

Owing to the low solubility of hydrocarbons in water, bacterial degradation takes place mainly at hydrocarbon-water interfaces. A. borkumensis SK2 attaches readily to the oil-water interface of hydrocarbon droplets in salt water (see > Fig. 3 and insert in > Fig. 4). The A. borkumensis SK2 genome has many determinants for the biosynthesis, export, modification and polymerization of exopolysaccharides, which are putatively involved in biofilm formation. Forty of these determinants are identified in a large region of lower G þ C content spread over almost 50 kb (position 1,011,000–1,058,000). This region codes for putative glycosyltransferases, as well as for proteins predicted to be involved in sugar nucleotide biosynthesis, the addition of non-carbohydrate decorations, export and polymerization of the polysaccharide constituents. A cluster putatively specifying alginate biosynthesis is located in another region (432,500– 448,500). The A. borkumensis SK2 chromosome hosts 16 pil genes related to those of Type IV pili, which mediate the formation of biofilms and are involved in surface translocations (Ochsner et al., 1994). A number of determinants for a Type II secretion system, Sec translocon and twin-arginine translocation (Tat) have also been identified (> Fig. 4). A number of genes related to secretion proteins were found, in particular, five genes putatively coding for HlyD family secretion proteins. Most of the above genes are likely to be involved in the formation of biofilms at oil-water and water-substratum interfaces. The 12 genes specifying biosynthesis of the alginate component of extracellular polysaccharides (EPS) in A. borkumensis SK2

borkumensis cell grown on hexadecane (courtesy of H. Lu¨nsdorf, GBF). The insert the right upper corner shows a TEM of A. borkumensis SK2 cells at the oil-water interface of hydrocarbon droplets in salt water. Predicted pathways for alkane degradation are depicted in marine blue. Predicted transporters are grouped by substrate specificity: inorganic cations (gray), inorganic anions (dark orange), amino acids/peptides/amines/purines/pyrimidines and other nitrogenous compounds (dark green), carboxylates (light green), drug efflux and other (dark gray). Export or import of solutes is designated by the direction of the arrow through the transporter. The energy coupling mechanisms of the transporters are also shown: solutes transported by channel proteins are shown with a double headed arrow; secondary transporters are shown with two arrowed lines indicating both the solute and the coupling ion; ATP-driven transporters are indicated by the ATP hydrolysis reaction; transporters with an unknown energy coupling mechanism are shown with only a single arrow. The P-type ATPases are shown with a double headed arrow to indicate they include both uptake and efflux systems. Where multiple homologous transporters with similar substrate predictions exist, the number of that type of protein is indicated in parentheses. Abbreviations: EPS extracellular polysaccharides; AA aminoacids; BCCT betaine/carnitine/choline transporters; GSP general secretion pathways; PRPP 5-phospho-alpha-D-ribose 1-diphosphate; Mhn complex sodium/proton antiporter involved in sodium excretion.

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are clustered. algD844KEIJFXLGA, (ABO_0384–0395) is located in a region (432,500– 448,500) that is predicted to be highly expressed (Reva et al., 2008). The gene algC, coding for phosphomannomutase (ABO_0937), is located within a large gene cluster harboring cell surface polysaccharide biosynthesis genes. The regulatory gene clusters algZR (ABO_2324, 2323) and algUSNMW (ABO_1639–1635) complete the alginate machinery set. Both the cluster organization, and the similarity at the amino acid level of the individual components, is akin to those of P. aeruginosa. Also the pathway for inner core region biosynthesis encoded by waaFP1CGP2 (ABO_2300–2296) is analogous to that in P. aeruginosa. This cluster neighbors rfaE (ABO_2290), which codes for the bifunctional protein for LPS biosynthesis and the gene lpxL1 (ABO_2292), which specifies lauryl acyltransferase involved in lipid A biosynthesis. However, there are no equivalents to the A- and B-band of the O-antigen structures of lipopolysaccharide LPS of P. aeruginosa (Rocchetta et al., 1999). Instead, a large cluster spread over almost 50 kb and comprising 40 genes typical for cell surface polysaccharide biosynthesis, was identified (ABO_0903 – ABO_0943). Most of the genes are predicted to be involved in sugar nucleotide biosynthesis (13 genes) and the addition of non-carbohydrate decorations (6 genes). There are also 10 putative glycosyltransferase genes, forming two distinct subgroups within the cluster, as well as genes likely to encode for proteins related to the export and polymerization of the polysaccharide constituents. Key constituents of a translocase/ polymerase-type export system were identified, including the genes wzb (ABO_0906), wzc (ABO_0907) and wzx (ABO_0915). Although this type of export machinery is known to control the production of complex heteropolymeric polysaccharides (Reeves et al., 1996), it was not possible to predict from the sequence data which kind of polysaccharide is encoded. Most genes for the murein sacculus are located in the clusters ABO_0591–0604 and ABO_1947–1959, whereas the rest of the genes is spread over the genome. The lipid A determinants lpxDAB are clustered within the ABO_1151–1156 region, together with genes involved in phospholipid biosynthesis, genes coding for an outer membrane proteins of the bacterial surface antigen family and ompH. This gene arrangement is conserved in P. aeruginosa, P. putida, E. coli, C. crescentus, S. meliloti, and I. loihiensis and reflects the fact that the lipid A part of the LPS is conserved within the Proteobacteria. Extensive transposon mutagenesis of the A. borkumensis genome helped to experimentally uncover a number of further features related to biofilm formation (Sabirova et al., 2008). For instance, the authors showed that development of a biofilm in Alcanivorax is likely to require autolysis, as witnessed by two specific biofilm-deficient mutants. Autolysis had been previously shown to provide ecological advantages during the development and dispersal of biofilms for another marine bacteria. Interestingly as well, Sabirova et al. also found that nitrogen metabolism seems to play an important role in biofilm formation by Alcanivorax as witnessed by the mutation in Abo_2302 encoding glnE glutamateammonia ligase adenylyltransferase. GlnE is a regulatory protein that is involved in the regulation of glutamine synthetase activity, which in turn is a key enzyme in nitrogen metabolism, which is responsible for the incorporation of ammonium into glutamate to produce glutamine at low ammonia concentrations. Sabirova et al. (2008) also found that the mini-Tn5 mutation in ABO_2248 encoding a putative ubiquinone biosynthesis protein was found to cause biofilm deficiency. This is consistent with the common observation that the respiratory status of cells is of relevance when growing as a biofilm. Ubiquinone is a key component of electron transport chain and a knockout mutation of the homologous ubiquinone biosynthetic gene in E. coli in an independent experiment (Søballe and Poole, 1998) resulted not only in the inability to

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synthesize ubiquinone but also severely diminished growth yields under aerobic conditions). These results hint at an important role of the quinone pool with respect to biofilm growth. Interestingly, in their functional analysis study, the authors have not detected any of the genes that might be involved in biofilm formation, as deduced earlier from the genome sequence (Schneiker et al., 2006), such as type-IV pili and type-II secretion proteins. This may indicate that the corresponding pathways in this organism can be redundant.

6

Nutrient Transport, Regulation, and Stress Responses

6.1

Nutrient Transport and Sodium Dependency

The success of a bacterium in the generally oligotrophic marine environment depends on effective scavenging of elements such as N, P, and S, and various oligo-elements such as Fe, Zn, Co, Mg, Mn, and Mo. A. borkumensis SK2 encodes genes for a broad range of transport proteins (> Fig. 4). These comprise determinants for about 50 permeases, of which roughly half belong to high affinity ABC-transport systems, five major facilitator superfamily (MFS) transport systems and two TRAP C4-dicarboxylate transporters (dctP, ABO_2146, and ABO_0688). No other carbohydrate transporter that are usually present in other bacteria were identified, which is consistent with its reported inability to grow on or to use many common, simple carbohydrates such as glucose (Yakimov et al., 1998). Consistent with its marine lifestyle, many of the transport systems in A. borkumensis SK2 are linked to Naþ pumps (> Fig. 4). The A. borkumesis SK2 genome encodes six subunits of a Naþ-dependent NADHquinone dehydrogenase (nqrABCDEF, ABO_1032–1037), enabling it to use the sodium gradient as a source of energy for nutrient uptake, taking advantage of the high salinity of the marine environment. Export of sodium ions at the expense of the proton gradient is performed by a variety of sodium/proton antiporters, including the multisubunit Naþ/Hþ antiporter (mnhABCDEFG genes, ABO_2653–2658), as well as NhaD (ABO_0238), NhaB (ABO_1226), NhaP (ABO_1404) and NhaC (ABO_2329) (> Fig. 4). A. borkumensis SK2 encodes also a symporter for Naþ/alanine (dagA, ABO_0618), Naþ/sulphate (ABO_0929), Naþ/glutamate (ABO_1478 and ABO_1616, see above) as well as several other Naþ-dependent symporters (ABO_327, ABO_1913, ABO_2155, ABO_2158, ABO_2490). The A. borkumensis SK2 genome encodes diverse and alternative systems for the uptake of both Dissolved Inorganic and Organic Nitrogen (Berman and Bronk, 2003). The DIN gene sets include two clusters of genes for active nitrate uptake and reduction (narKGHIJ, respectively, ABO_0547–0543, and nrtCB-nasDTS, ABO_0851–0860). Interestingly, the nar cluster is flanked by ptsHN genes, which code for a NPr-related protein and a nitrogen regulatory IIA protein of a phosphotransferase system, respectively and by the rpoN (RNA-polymerase s54 factor) and rpoX (s54 modulation protein) genes. Like many nitrogen-related functions, nitrate assimilation and transport are known to be s54 dependent in pseudomonads and other g-Proteobacteria (Cases et al., 2003). To complete the DIN gene set, the A. borkumesis SK2 genome encodes as well for three high-affinity ammonium transporter systems (amt), of which two (ABO_1124 and ABO_2529, amtB homologues) are immediately flanked by one nitrogen regulator gene (glnK) each, whereas the third (ABO_1476) is clustered with determinants for GltP, a sodium/dicarboxylate symporter, a short-chain dehydrogenase/reductase and a response regulator (Rrp1). In contrast, versatile, ubiquitous soil bacteria such as

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P. aeruginosa and P. putida, with genomes twice as large as A. borkumensis SK2, have only two ammonium transporters. One of these is flanked by a nitrogen regulator. In P. putida and E. coli, the regulation of the glnK-amtB operon itself is linked to that of ntrC in response to nitrogen availability. ntrCB genes, ABO_2259,2260, code for a two-component sensor regulator and its cognate histidine kinase in A. borkumensis SK2. Regulation of both the glnK-amtB and glnA-ntrCB operons are s54 dependent (Cases et al., 2003). Possibly, the multiple transporters may have different affinities and may be regulated differently depending on the nutrient concentration. The inter-twining of these various layers of regulation and the diversity of transport, assimilation and control mechanisms in A. borkumensis, reflects the emphasis it puts on effective and efficient nitrogen scavenging from diluted environments. This appears to be a signature of aquatic, nutrient-poor habitats, as such diversity and profusion of nitrogen uptake and assimilation mechanisms is relatively widespread in a number of other marine or fresh-water microorganisms (e.g., Synechocystis, Synechococcus, Prochlorococcus, Idiomarina, Nostoc, Chlorobium, etc.). The DON gene sets pertain one operon for uptake and assimilation of urea (ureABCDEFGJ, ABO_2715–2722), two for branched-chain amino acid transporters (ABO_1982–1985 and ABO_2509–2512) and one for active oligopeptide transport (Opp, ABO_1216–1220, > Fig. 4). Phosphate uptake is mediated by a typical, high-affinity ABC-type Pst system coded phoU-pstBACS, ABO_2681–2685 and phoBR (ABO_0166, 0167), as well as by a lowaffinity transporter protein (coded by ABO_2305). For sulphate uptake and in addition to three putative MFS permeases (ABO_1512, ABO_1943, and ABO_1945), A. borkumensis SK2 codes for a homologue of CysZ, another putative permease (ABO_0946) and putative sodium/ sulphate symporter (ABO_0929). An ABC-sulfonate uptake system is encoded by ABO_1887– 1889. Phosphate uptake is mediated by a typical, high-affinity ABC-type Pst system coded phoU-pstBACS, ABO_2681–2685 (clustered with arsCR for arsenic/arsenate resistance) and phoBR (ABO_0166, 0167), as well as by a low-affinity transporter protein (coded by ABO_2305). In addition to this array of N, P, and S uptake systems, A. borkumensis SK2 specifies the import of various oligo-elements, such as molybdate (modABC, ABO_1254– 1256), zinc (znuAB, ABO_0155–0157), cobalt (ABO_1335), cobalamin (ABO_2368–2370) and magnesium (mgtE, ABO_0541). This wealth of scavenging abilities enables A. borkumensis SK2 to exploit its alkanedegrading potential to tackle quickly and effectively the sudden increase in carbon availability and resulting carbon:nutrient imbalance that ensues from typical oil-spills, and may well provide A. borkumensis SK2 with a significant competitive advantage over other prevalent hydrocarbon degraders. Also, the presence of three rRNA operons in A. borkumensis SK2 is consistent with its potential to respond swiftly to changes in resource availability (Klappenbach et al., 2000).

6.2

Stress Responses and Osmoregulation

In addition to the common stress response systems, the A. borkumensis SK2 genome specifies a number of habitat-related responses. Like the genome of the marine bacterium I. loihiensis, that of A. borkumensis SK2 encodes three Naþ-driven multidrug efflux pumps (ABO_0158, ABO_1554, and ABO_2623), as well as several systems for detoxification of arsenate and salts of mercury, copper and other heavy metals (> Fig. 4). Halotolerant bacteria generally accumulate Kþ, glutamate, and the compatible solutes ectoine and betaine as osmoprotectants.

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In addition to uptake systems for glutamate (GltP, ABO_1478, GltS, ABO_1616) and choline/ betaine (ABO_0232, ABO_0637, and ABO_0808), the A. borkumensis SK2 genome contains genetic determinants for biosynthesis of glutamate (gltA, ABO_1501, gltB, ABO_2229, gltD, ABO_2228 and gltX, ABO_1509), ectoine (ectABC, ABO_2150_2152) and betaine (ABO_0886) (> Fig. 4). However, genes for the synthesis of other organic compatible solutes, such as trehalose, di-inositol phosphate or mannosylglycerate, were not identified. Moreover, as is the case for the genome of I. loihiensis, determinants for ATP-dependent Kþ transporters are absent, which may indicate a strategy for minimization of ATP expenditure in response to consistently high salt concentrations in marine environments. Some of these hypotheses drawn from the analysis of genome sequence were latter confirmed through multitransposon mutagenesis study by Sabirova et al. (2008), who showed, for instance, that one of the osmosensitive A. borkumensis mini-Tn5 mutants was deficient in the ectB gene, encoding 2,4-diaminobutyrate aminotransferase, a key enzyme of the biosynthesis pathway of ectoine. Not unexpectedly, the analysis of A. borkumensis osmosensitive Tn5 mutants has also pointed at several efflux pumps that evidently are essential for this organism’s ability for osmoadaptation. Among these were the ABC export system (ABO_0250), the ABC efflux transporter (ABO_1673), and the so-called multidrug and toxin extrusion (MATE) efflux family protein (ABO_2623). Moreover, mRNA decay and modification also seem to play a crucial role in adaptation to salt, as a mutation in the gene ABO_0333 encoding polynucleotide phosphorylase was found to cause osmosensitivity. Polynucleotide phosphorylase was found to add heteropolymeric tails to the 30-ends of RNA transcripts (Mohanty and Kushner, 2003), which could lead to selective accumulation of osmotic shock-characteristic mRNA transcripts and/or posttranscriptional modification of mRNAs. Repair of UV-induced DNA damage requires a robust coordination of the expression of DNA repair genes. Sabirova et al. (2008), found a number of UV-sensitive A. borkumensis mutants (deficient in genes ABO_0427, ABO_0945, ABO_1305, ABO_1801, ABO_0753, ABO_0249, and ABO_1474, respectively) that were of affected in their function in DNA repair. These genes can be grouped into three subgroups based on their respective molecular functions, namely (ABC)-excision repair, SOS-response, and mismatch repair. All these seem to be crucial mechanisms involved in the repair of DNA damages caused by UV stress in Alcanivorax. In addition to the cellular nucleic acid metabolism being affected by UV, A. borkumensis also developed specific strategies to protect itself against oxidative stress resulting from UV radiation, as evidenced by several of the UV sensitive mini-Tn5 mutants that were found to be deficient in certain components of the respiratory chain, like nitrate reductase, cytochrome b561, and iron–sulfur proteins. As shown by Sabirova et al. (2008), adaptation to low-temperature conditions in A. borkumensis seems to proceed via various regulatory mechanisms and these regulatory systems are not overlapping with the ones of the other tested phenotypes, meaning that adaptation to low temperature proceeds via independent regulatory mechanisms. For instance, one key regulatory element important for this organism’s ability to adapt to low temperatures is the BipA regulator encoded by ABO_2269, which functions as a translation factor required specifically for expression of the transcriptional modulator Fis. Expression of BipA triggers efficient induction of the fis gene, thereby in turn modulating the expression of a range of Fis-dependent downstream processes, including coping with a temperature downshift (Kim et al., 2004). Other mechanisms include adaptation of the fluidity of the cell membrane through regulation of the synthesis of lipids containing branched-chain fatty acids; the modulation of stress-response regulators such as those from the MerR family; and

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the regulation of control of intracellular levels of glutamine and glutamate, presumably acting under these conditions as cryoprotectors. Altogether, the detailed experimental study by Sabirova et al. revealed that adaptation of A. borkumensis to UV radiation, low temperature, osmotic stress, and biofilm formation in each case involves a multiplicity of diverse cellular functions contributing to the respective protective responses. Furthermore, the authors found that A. borkumensis is likely to use the protection mechanisms being attributed so far only to eukaryotic organisms (cold-induced posttranslational modification by methylation), thus enriching its repertoire of molecular mechanisms to fight against the environmental stress.

6.3

Regulation and Signal Transduction in A. borkumensis SK2

Analysis of the A. borkumensis SK2 genome sequence reveals 87 genes putatively coding for transcriptional regulators of various families: including 17 AraC determinants, which are generally involved in the regulation of a variety of processes such as carbon metabolism and stress responses (e.g., MmsR, coded by ABO_0019, involved in valine pathway); 16 TetR, which typically regulate expression of resistance to hydrophobic detergents and antibiotics, as well as polymer metabolism (e.g., PhaD (ABO_0088) regulator of polyalkanoate metabolism); 15 LysR, which are associated to a wide range of processes including central metabolism and stress responses (e.g., OxyR, key regulator of oxidative response network, coded by ABO_0169); 6 MerR (e.g., ABO_0765 putatively regulating photolyase coded by ABO_0766); 5 ArsR (3 of which regulate arsenic resistance operons as described above); 4 LuxR (e.g., AlkS, coded by ABO_2706, the positive regulator of the alk operon); 4 GntR (e.g., the product of gene ABO_0121 that regulates the expression of the second alkane monoxygenase gene, alkB2, ABO_0122) and 3 MarR (e.g., ABO_2149, regulating a TRAP dicarboxylate transporter); as well as common Fur, Anr, CobB, PyrR and single members of the Fis, Crp, PadR family, among others. This global number of transcription regulator genes (3.2% of all CDS in A. borkumensis SK2) is in line with the generally observed trend of an increasing proportion of regulators with genome size (rate of roughly 1% ORFs per megapase pairs (Lombardot et al., 2005)). The A. borkumensis SK2 genome specifies also for 25 putative histidine kinases (of which four are hybrid) and 29 response regulators. The numbers are similar to those of I. loihiensis (comparable genome size) and about four times fewer than those of the soil bacteria P. putida and P. aeruginosa (genomes twice as large). Seven of the A. borkumensis SK2 encoded response regulators consist of a CheY-like receiver domain and a winged-helix DNA-binding domain. 11 proteins contain a GGDEF output domain and one contains an EAL domain. Both domain families have been shown to be often linked to the production of EPS (Galperin et al., 2001) and are likely to play a similar role in A. borkumensis SK2. The number of sigma factors specified by a genome – in particular those of the s70 of the ECF-subfamily, which increase more than proportionally with genome size, reflect the ability of an organism to build adaptive responses to environmental signals (Martı´nez-Bueno et al., 2002; Cases et al., 2003). In line with this, the genome of A. borkumensis SK2 specifies nine sigma factors of which three belong to the ECF-subfamily (AlgU encoded by ABO_1639, RpoE by ABO_2179 and a predicted transcriptional regulator by ABO_2175). In comparison, the versatile P. putida and P. aeruginosa, which thrive in more diverse and changing environments, specify 24 sigma factors, of which 19 belong to the ECF-subfamily. Also, the presence of

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three rRNA operons in A. borkumensis SK2 is consistent with its potential to respond swiftly to changes in resource availability (Klappenbach et al., 2000). The functional analysis study of the A. borkumensis’s stress response by means of transposon mutagenesis carried out by Sabirova et al. (2008) complemented experimentally the above genome sequence analysis and revealed a broad overlap between the UV, salt-, and biofilm-deficient mutants phenotypes study, and exclusively on the level of signal transduction and regulation. For instance, the authors found that biofilm formation and adaptation to UV and salt appear to require a two-component hybrid sensor regulator protein encoded by ABO_1986 and ABO_1987 (interrupted by mini-Tn5 in one UV mutant, in one osmosensitive mutant, and in one biofilm-deficient mutant). ABO_1987 shows the typical domain arrangement of histidine kinases and, together with the upstream gene (ABO_1986), comprises a two-component hybrid sensor regulator protein, which is highly conserved across marine bacterial species. UV adaptation and biofilm formation appear also to be linked via another signal transduction system, a conserved hypothetical signal transduction protein, encoded by ABO_2433 and found to be interrupted in one UV mutant and one biofilm mutant. The regulator encoded by ABO_2433 displays a complex domain arrangement with three PAC, three PAS, one GGDEF, and one EAL domains. PAS domains are important signaling modules that monitor changes in light intensity, redox potential, oxygen tension, and/or the overall cellular energy level. GGDEF and EAL domains are known to be also involved in the metabolism of the secondary messenger cyclo-diguanylate (c-di-GMP). c-di-GMP had been found to play a key role in biofilm formation in other bacteria by stimulating cell aggregation and surface attachment (Tischler and Camilli, 2004). Thus, in A. borkumensis the cellular stress response to UV stress and biofilm formation may be triggered by changes in the intracellular oxygen and c-di-GMP levels. These and other experimental findings by Sabirova et al. (2008) show that there are numerous cross connections between different phenotypes (e.g., biofilm and UV stress; biofilm and UV and osmoadaptation) on signal transduction level, confirming the hypotheses derived from the genome analysis above (see also Schneiker et al., 2006) that complex and tightly controlled cellular interactions involving oxygen as a primary messenger and cyclic-di-GMP as a secondary messenger are required for Alcanivorax responses to environmental stresses. Alcanivorax seems to use overlapping signal transduction systems linking biofilm formation, UV tolerance, and osmoadaptation. As a whole these results provide valuable insights into bacterial function in a complex marine environment.

7

Metabolic Specialization in Oligotrophic Environments

A. borkumensis SK2, like other hydrocarbonoclastic bacteria, can grow on a very restricted range of substrates (Yakimov et al., 1998). The absence of a functional PEP-dependent sugar/phosphotransferase system or other type of sugar transporter, as well as the lack of several determinants for key enzymes of the glycolytic, pentose phosphate shunt and Entner– Doudoroff pathways, is consistent with the demonstrated inability of A. borkumesis SK2 to utilize simple hexoses and other simple carbohydrates for growth. Examples of missing key enzymes include glucokinase (Glk), glucose-6-phosphate dehydrogenase (Zwf), aldose epimerase (GalM), 2-keto-3-deoxy-6-phosphogluconate aldolase (Eda) and phosphogluconate dehydratase (Edd). However, it codes for many of the enzymes required for glycolysis/

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gluconeogenesis, and has a full complement of candidate genes for the citric acid cycle with glyoxylate shunt (> Fig. 4). ATP synthesis is mediated by an F-type ATP synthase (ABO_2725–2733). Several central enzymes of sugar metabolism, including transketolase (TktA, ABO_2615), ribose 5-phosphate isomerase (RpiA, ABO_2606) and ribulose-5phosphate-3-isomerase (ABO_2042) are encoded by the A. borkumensis SK2 genome and appear to be sufficient to provide intermediates for biosynthesis of nucleotides, several cofactors and amino acids. Genes for the synthesis of all 20 amino acids from chorismate, pyruvate, 3-phosphoglycerate, glutamate, and oxaloacetate are present, as are those of pathways for the synthesis of purines, pyrimidines, and nucleotides. Analysis of the genome suggests also that A. borkumensis SK2 is capable of both synthesis and elongation of fatty acids from pyruvate and hydrocarbons, a prediction that has been confirmed experimentally (Sabirova et al., 2006b). Genes for a b-oxidation pathway are present, as are determinants for the synthesis of an extensive range of cofactors and prosthetic groups, including biotin, folic acid, ubiquinone, pantothenate, coenzyme A, ubiquinone, glutathione, thioredoxin, riboflavin, FMN, FAD, NAD, NADP, porphyrin, thiamin, cobalamin, pyridoxal 5’-phosphate, tetrahydrofolate, and lipoate.

7.1

Glyoxylate Bypass and Gluconeogenesis

During growth on alkanes as the sole carbon source, bacteria must generate all cellular precursor metabolites from acetyl-CoA, the main intermediate formed during alkane degradation via b-oxidation of fatty acids (> Fig. 5). One mechanism Alcanivorax utilizes is the short circuiting of the citric acid cycle, through up-regulation of the glyoxylate bypass, which routes acetyl-CoA to the key 3-carbon metabolite phosphoenolpyruvate, via isocitrate, glyoxylate, and malate, by means of isocitrate lyase and malate synthase, thereby reducing the activity of the CO2-releasing steps of the: alkane-grown cells exhibit up-regulation of ABO_2741, encoding isocitrate lyase AceA, and ABO_1267, encoding malate synthase GlcB, and down-regulation of enzymes mediating CO2-releasing steps of the tricarboxylic acid (TCA) cycle short circuited by the glyoxylate shunt, including ABO_1281, encoding isocitrate dehydrogenase Icd, and ABO_1494, encoding 2-oxoglutarate dehydrogenase LpdG. We also suspect down-regulation of another enzyme indicative of a complete TCA cycle, namely, succinyl-CoA synthetase SucC, encoded by ABO_1493, as according to our in silico analysis, this gene is located in the same putative operon as ABO_1494. Another enzyme of the TCA cycle needed for the glyoxylate bypass, namely, succinate dehydrogenase SdhD (ABO_1499), was also found to be up-regulated in alkane-grown cells. The hypothesis that all biosynthetic precursors derive from acetyl-CoA in alkane-grown cells is also consistent with the finding that enzymes involved in gluconeogenesis, namely, malic enzyme MaeB (ABO_2239) and phosphoenolpyruvate synthase PspA-1 (ABO_1427), were up-regulated. Thus, the key metabolic intermediate in alkane-grown cells is malate, formed through the channeling of acetyl-CoA into the glyoxylate bypass. The above analysis of the metabolic, transport and regulatory features of A. borkumensis reveals much about its ecological survival strategies. In fact, although it best thrives, degrades and competes is in the presence of large amounts of oil, the more frequent conditions that A. borkumensis and OHCB‘s in general must face are waters with very low concentrations of hydrocarbons, which requires a sharply adapted metabolism. The genome of A. borkumensis shows that it is well adapted to oligotrophic conditions as:

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. Figure 5 Alkane growth-induced up-regulation of the glyoxylate bypass and gluconeogenesis. The glyoxylate bypass is carried out by isocitrate lyase (1) and malate synthase (2). Succinate produced via glyoxylate bypass is converted to malate by succinate dehydrogenase (3). Malate is converted to either oxaloacetate by malate dehydrogenase (4), or is used by malic enzyme (5) in gluconeogenesis to produce pyruvate. Pyruvate is then converted by phosphoenolpyruvate synthase (6) to produce phosphoenolpyruvate. Up-regulation of the glyoxylate shunt is accompanied by a down-regulation of 2-oxoglutarate dehydrogenase. Adapted from Sabirova et al. (2006a, b).

 it has a relative large number of determinants for specific transporters fostering both substrate and nutrient uptake;

 it has, along with ‘‘eutrophic’’ (inducible under elevated concentration) transporters, a good number of more ‘‘oligotrophic’’ transporters for N, S, and P;

 its cytoplasm is typically very diluted (very few proteins expressed) and, importantly, all

essential enzymes (e.g., alk genes) that are involved in the ‘‘launch’’ metabolism are inducible.

This adaptability to both eutrophic and oligotrophic lifestyles is a striking feature of A. borumensis (and likely of OHCB’s in general). The combination of R-and K-strategies likely endows OHCB’s with an important competitive advantage.

8

Genome-Scale, Constraint-Based Modeling of the A. borkumensis Metabolism

The relationship between the genotype and the phenotype is complex, highly nonlinear and cannot be predicted from simply cataloguing and assigning gene functions to genes found in the genome, as described hitherto. Comprehensive understanding of cellular metabolism

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requires placing the function of every gene in the context of its role in attaining the set goals of a cellular function. This demands the integrated consideration of many interacting components, which is best done through mathematical modeling. Hence, drawing on annotated genome sequence data, biochemical information and strain-specific knowledge, we developed a genome-wide, quantitative framework based on constraint-based modeling of metabolic and transport network of A. borkumensis SK2 (for methods description see Puchalka et al., See > Chapter 75, Vol. 5, Part 5). The model, currently comprising a network of 462 metabolic genes associated with 490 reactions and 478 distinct metabolites, provides a framework to study the consequences of alterations in the genotype and to gain insight into the phenotype– genotype relation. The model was experimentally validated by comparison of predictions with measurements described in Sabirova et al. (2006b) for a number of physiological parameters. Using this modeling framework, we then explored the metabolic space of A. borkumensis SK2 to identify structures in the networks that may lead to enhanced production of polyhydroxyalkanoates (PHAs) of biotechnological value.

8.1

Storage Compounds and Polyhydroxyalkanoate Production by A. borkumensis

For carbon-limited microbes, an increase in carbon allows an increase in growth rate until another growth limitation is reached. Under conditions of high C/N ratios, many microbes synthesize carbon storage materials, like PHAs, which enable them to survive during less nutritious periods. The appearance of alkanes in oligotrophic environments, for instance, following an oil-spill in the ocean, allows Alcanivorax to ‘‘bloom’’ initially until nitrogen limitation is experienced, after which it accumulates storage compounds (Abraham et al., 1998). Sabirova et al. (2006a, b) has shown that A. borkumensis produces either mediumchain-length PHAs, consisting of 3-hydroyacyl monomers of 6–12 carbon units, or polyhydroxybutyrate (PHB) (four carbon units), depending on the carbon source used for growth. Alcanivorax ABO_1418, one of two A. borkumensis phaC PHA synthase genes, was solely expressed in alkane-grown cells, whereas another, ABO_2214, was not expressed at detectable levels in such cells. Since PHA is also produced at high C/N ratios in cells grown on pyruvate (Sabirova et al., 2006b), it seems that A. borkumensis produces different PhaC PHA synthases in response to different growth substrates; these distinct enzymes may have different specificities that reflect distinct metabolites produced from the different growth substrates. However, although A. borkumensis is able to produce these various PHAs under different conditions, it does so in less amounts, in absolute terms, than other more ‘‘specialized’’ organisms. Recent work (Capello & Yakimov, > Chapter 6, Vol. 3, Part 1) has shown that acylglycerols (TAGs) and wax esters (WEs) are the predominant hydrophobic storage compounds occurring in this strain. A peculiarity of storage lipids from A. borkumensis, compared to those from other bacteria, is their relatively low content of unsaturated fatty acids. In particular, WEs and TAGs produced during cultivation on hexadecane almost exclusively contained saturated fatty acids. Using the constraint-based model framework, we assessed the quantitative influence of carbon, nitrogen and phosphorus limitation over the growth of A. borkumensis, and found that biomass and PHB formation were clearly more affected than respiration process by the limitation on either, nitrogen or phosphorus sources. Thus, as expected, nitrogen and phosphorus are critical components for biomass and PHB formation in A. borkumensis when the

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alkanes are available in non-limiting amounts. On the contrary, the aerobic respiration process was clearly more affected by carbon limitation since the oxygen consumption and the carbon dioxide formation showed significant low values compared with the other two limitedconditions. Similar conditions were imposed to E. coli to experimentally study its growth parameters under carbon and nitrogen limited conditions (Sauer et al., 1999) and, regarding the biomass formation and the respiration process, its results agreed with those found for A. borkumensis (Garcia et al., unpublished). The model described well the differences in PHB formation by A. borkumensis bacteria growing on pyruvate and octadecane, namely, that the amount of PHB produced from alkane (i.e., under conditions of a high C/N ratio) was three times higher than that produced upon growth on pyruvate. This significant difference is due to the different pathways to PHB depending on the carbon source. During growth on pyruvate as carbon source, PHB is formed from acetyl-CoA that was further converted to acetoacetyl-CoA and (S)-3Hydroxybutyryl-CoA, as precursor of ePHB formation by the enzymes acetyl-CoA acyltransferase and (S)-3-Hydroxybutyryl-CoA dehydrogenase, respectively. When A. borkumensis grows on octadecane as a carbon source, PHB is generated via alkane oxidation and fatty acid metabolism in the PHB precursor, (S)-3-Hydroxybutyryl-CoA. Finally, the model confirmed the notable increment in PHA formation that was achieved when the enzyme ‘‘tesB-like’’ (a hydroxyacyl-CoA-specific thioesterase acting exclusively on hydroxylated acyl-CoAs), was constrained to zero thus mimicking an A. borkumensis tesB-like mutant. As demonstrated experimentally by Sabirova et al. (2006b), blockage of this enzyme results in the rechanneling of CoA-activated hydroxylated fatty acids, the cellular intermediates of alkane degradation, towards PHA (> Fig. 6). The formation of PHA in the tesB-like mutant growing on alkane was 20 times higher than in WT strain under the same growth conditions. Surprisingly, and contrary to most other bacteria, these compounds in the A. borkumensis mutant are excreted and accumulate extracellularly, which is of great biotechnological value. In summary, an experimentally-validated genome-scale metabolic model assists the process of hypothesis generation and to gain insight into the basis of hydrocarbonoclastic, marine lifestyle of A. borkumensis, its genomic responses to environmental stresses, the ability to degrade a range of hydrocarbons and to dominate oil-degrading microbial communities, as well on the mechanisms that provide it with its remarkable oil-degrading abilities and its competitive advantage in oil-polluted environments. The modeling of alkane fluxes versus those of nitrogen and phosphorus through the metabolic network also allowed the discovery of conditions in which the excess carbon available in hydrocarbons was not directly translated into bacterial biomass. Instead of that, carbon overflow was diverted to the production of polyhydroxyalkanoates, an activity for which A. borkumensis SK2 showed to be genetically well endowed.

9

Concluding Remarks and Research Needs

The analysis of the genome sequence and functional studies of A. borkumensis SK2 has provided significant new insights into the genomic basis of (1) its unusual metabolic capability: a highly restricted range of growth substrates but high catabolic capacity for diverse hydrocarbon substrates, (2) its effectiveness at accessing the hydrophobic and poorly bioavailable oil hydrocarbons, (3) its efficiency in accessing other nutrients and micronutrients,

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. Figure 6 Suggested pathway of PHA biosynthesis in A. borkumensis SK2 and mutant strain C9 grown on hydrocarbons. Hydrocarbons are degraded via terminal oxidation to produce free fatty acids, which are then activated by an acyl-CoA synthase and subjected to b-oxidation. The (S)-3-OH-acyl-CoAs produced by b-oxidation are isomerized into (R)-3-OH-acyl-CoAs by the action of an isomerase. (R)-3-OH-Acyl-CoAs produced during b-oxidation are converted to either 3-HAA through the action of TesB-like acyl-CoA thioesterase or PHA through the action of PhaC synthase. The mutation in the TesB-like acyl-CoA thioesterase abolishes the production of free 3-HAA and channels (R)-3-OH-acyl-CoAs. From Sabvirova et al. (2006b).

(4) its ability to overcome carbon:nutrient imbalances typical of oil-spills, and (5) its tolerance of the stresses it faces in its natural environment. The genomic basis of these features underlies at least part of the ecological success of A. borkumensis and its environmental importance in the natural biodegradation of oil hydrocarbons entering marine systems. These insights should aid the development of new strategies to accelerate the growth and potentiate the catabolic activities of Alcanivorax immediately following an oil spill, and thus to develop improved ecological damage mitigation measures. The genomic analysis has also revealed other potential biotechnological applications, namely the hyper-production of PHA and its release from the cell, a thus far unique phenotype in PHA biology which raises the interesting possibility of extracellular polyhydroxyalkanoates for the production of bioplastics (Sabirova et al., 2006b). Another category of application is biocatalysis, as the A. borkumensis SK2 genome encodes a number of proteins putatively involved in metabolic reactions of potential biotechnological interest, including eight hydrolases of the haloacid dehydrogenase/epoxide

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family, two determinants for dienelactone hydrolases (ABO_1618 and ABO_1886), three for deacylases, 36 for various cytochrome proteins and 30 for oxidoreductases of different families and with unknown substrate esters. Various oxidoreductase genes are clustered together and/ or located in operon-like structures with determinants for monoxygenases, aldehyde dehydrogenases, decarboxylases, Rieske 2Fe–2S family proteins, etc. The genome also includes 11 genes coding for different lipases/esterases of unknown specificity. Two of these esterases were purified and functionally characterized: they exhibit excellent enzymatic activity (up to two orders of magnitude higher than typical esterases), have wide substrate profiles, remarkable enantio-selectivities and chemical tolerance, and may thus have potential for the resolution of chiral mixtures in biocatalysis (Ferrer et al., unpublished). Thus, clearly, a major research need is the detailed functional analysis and characterization of the wealth of, as yet, uncharacterized genes and gene products of A. borkumensis. Only then, can the full potential of this fascinating bacterium be exploited for biotechnological and environmental applications.

References Abraham WR, Meyer H, Yakimov M (1998) Novel glycine containing glucolipids from the alkane using bacterium Alcanivorax borkumensis. Biochim Biophys Acta 1393: 57–62. Berman T, Bronk DA (2003) Dissolved organic nitrogen: a dynamic participant in aquatic ecosystems. Aquat Microb Ecol 31: 279–305. Bruns A, Berthe-Corti L (1999) Fundibacter jadensis gen. nov., sp. nov., a new slightly halophilic bacterium, isolated from intertidal sediment. Int J Syst Bacteriol 49: 441–448. Cappello S, Denaro R, Genovese M, Giuliano L, Yakimov MM (2007) Predominant growth of Alcanivorax during experiments on ‘‘oil spill bioremediation’’ in mesocosms. Microbiol Res 162: 185–190. Cases I, De Lorenzo V, Ouzounis CA (2003) Transcription regulation and environmental adaptation in bacteria. Trends Microbiol 11: 248–253. Ferna´ndez-Martı´nez J, Pujalte MJ, Garcı´a-Martı´nez J, Mata M, Garay E, Rodrı´guez-Valera F (2003) Description of Alcanivorax venustensis sp. nov. and reclassification of Fundibacter jadensis DSM 12178T (Bruns and Berthe-Corti 1999) as Alcanivorax jadensis comb. nov., members of the emended genus Alcanivorax. Int J Syst Evol Microbiol 53: 331–338. Galperin MY, Nikolskaya AN, Koonin EV (2001) Novel domains of the prokaryotic two-component signal transduction systems. FEMS Microbiol Lett 203: 11–21. Harayama S, Kishira H, Kasai Y, Shutsubo K (1999) Petroleum biodegradation in marine environments. J Mol Microbiol Biotechnol 1: 63–70. Kalscheuer R, Sto¨veken T, Malkus U, Reichelt R, Golyshin PN, Sabirova JS, Ferrer M, Timmis KN,

Steinbu¨chel A (2007) Analysis of storage lipid accumulation in Alcanivorax borkumensis: evidence for alternative triacylglycerol biosynthesis routes in bacteria. J Bacteriol 189: 918–928. Kasai Y, Kishira H, Syutsubo K, Harayama S (2001) Molecular detection of marine bacterial populations on beaches contaminated by the Nakhodka tanker oil-spill accident. Environ Microbiol 3: 246–255. Kasai Y, Kishira H, Sasaki T, Syutsubo K, Watanabe K, Harayama S (2002) Predominant growth of Alcanivorax strains in oil-contaminated and nutrient-supplemented sea water. Environ Microbiol 4: 141–147. Kim BH, Kim HG, Bae GI, Bang IS, Bang SH, Choi JH, Park YK (2004) Expression of cspH upon nutrient up-shift in Salmonella enterica serovar Typhimurium. Arch Microbiol 182: 37–43. Klappenbach JA, Dunbar JM, Schmidt TM (2000) rRNA operon copy number reflects ecological strategies of bacteria. Appl Environ Microbiol 66: 1328–1333. Liu C, Shao Z (2005) Alcanivorax dieselolei sp. nov., a novel alkane-degrading bacterium isolated from sea water and deep-sea sediment. Int J Syst Evol Microbiol 55: 1181–1186. Lombardot T, Bauer M, Teeling H, Amann R, Glo¨ckner FO (2005) The transcriptional regulator pool of the marine bacterium Rhodopirellula baltica SH 1T as revealed by whole genome comparisons. FEMS Microbiol Lett 242: 137–145. Martı´nez-Bueno MA, Tobes R, Rey M, Ramos JL (2002) Detection of multiple extracytoplasmic function (ECF) sigma factors in the genome of Pseudomonas putida KT2440 and their counterparts in Pseudomonas aeruginosa PA01. Environ Microbiol 4: 842–855.

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Mohanty BK, Kushner SR (2003) Genomic analysis in Escherichia coli demonstrates differential roles for polynucleotide phosphorylase and RNase II in mRNA abundance and decay. Mol Microbiol 50: 645–658. Ochsner UA, Fiechter A, Reiser J (1994) Isolation, characterization, and expression in Escherichia coli of the Pseudomonas aeruginosa rhlAB genes encoding a rhamnosyltransferase involved in rhamnolipid biosurfactant synthesis. J Biol Chem 269: 19787–19795. Reeves PR, Hobbs M, Valvano MA, Skurnik M, Whitfield C, Coplin D, Kido N, Klena J, Maskell D, Raetz CRH, Rick PD (1996) Bacterial polysaccharide synthesis and gene nomenclature. Trends Microbiol 4: 495–503. Reva ON, Hallin PF, Willenbrock H, Sicheritz-Ponten T, Tu¨mmler B, Ussery DW (2008) Global features of the Alcanivorax borkumensis SK2 genome. Environ Microbiol 10: 614–625. Rivas R, Garcı´a-Fraile P, Peix A, Mateos PF, Martı´nezMolina E, Vela´zquez E (2007) Alcanivorax balearicus sp. nov., isolated from Lake Martel. Int J Syst Evol Microbiol 57: 1331–1335. Rocchetta HL, Burrows LL, Lam JS (1999) Genetics of O-antigen biosynthesis in Pseudomonas aeruginosa. Microbiol Mol Biol Rev 63: 523–553. Ro¨ling WFM, Milner MG, Jones DM, Lee K, Daniel F, Swannell RJP, Head IM (2002) Robust hydrocarbon degradation and dynamics of bacterial communities during nutrient-enhanced oil spill bioremediation. Appl Environ Microbiol 68: 5537–5548. Ro¨ling WFM, Milner MG, Jones DM, Fratepietro F, Swannell RPJ, Daniel F, Head IM (2004) Bacterial community dynamics and hydrocarbon degradation during a field-scale evaluation of bioremediation on a mudflat beach contaminated with buried oil. Appl Environ Microbiol 70: 2603–2613. Ron EZ, Rosenberg E (2001) Natural roles of biosurfactants. Environ Microbiol 3: 229–236. Sabirova JS, Chernikova TN, Timmis KN, Golyshin PN (2008) Niche-specificity factors of a marine oildegrading bacterium Alcanivorax borkumensis SK2. FEMS Microbiol Lett 285: 89–96. Sabirova JS, Ferrer M, Regenhardt D, Timmis KN, Golyshin PN (2006a) Proteomic insights into metabolic adaptations in Alcanivorax borkumensis induced by alkane utilization. J Bacteriol 188: 3763–3773. Sabirova JS, Ferrer M, Lu¨nsdorf H, Wray V, Kalscheuer R, Steinbu¨chel A, Timmis KN, Golyshin PN (2006b) Mutation in a ‘‘tesB-like’’ hydroxyacyl-coenzyme

A-specific thioesterase gene causes hyperproduction of extracellular polyhydroxyalkanoates by Alcanivorax borkumensis SK2. J Bacteriol 188: 8452–8459. Sauer U, Lasko DR, Fiaux J, Hochuli M, Glaser R, Szyperski T, Wu¨thrich K, Bailey JE (1999). Metabolic flux ratio analysis of genetic and environmental modulations of Escherichia coli central carbon metabolism. J Bacteriol 181: 6679–6688. Schneiker S, Dos Santos VAPM, Bartels D, Bekel T, Brecht M, Buhrmester J, Chernikova TN, Denaro R, Ferrer M, Gertler C, Goesmann A, Golyshina OV, Kaminski F, Khachane AN, Lang S, Linke B, McHardy AC, Meyer F, Nechitaylo T, Pu¨hler A, Regenhardt D, Rupp O, Sabirova JS, Selbitschka W, Yakimov MM, Timmis KN, Vorho¨lter FJ, Weidner S, Kaiser O, Golyshin PN (2006) Genome sequence of the ubiquitous hydrocarbon-degrading marine bacterium Alcanivorax borkumensis. Nat Biotechnol 24: 997–1004. Søballe B, Poole RK (1998) Requirement for ubiquinone downstream of cytochrome(s) b in the oxygenterminated respiratory chains of Escherichia coli K-12 revealed using a null mutant allele of ubiCA. Microbiology 144: 361–373. Tischler AD, Camilli A (2004) Cyclic diguanylate (c-diGMP) regulates Vibrio cholerae biofilm formation. Mol Microbiol 53: 857–869. van Beilen JB, Marı´n MM, Smits THM, Ro¨thlisberger M, Franchini AG, Witholt B, Rojo F (2004) Characterization of two alkane hydroxylase genes from the marine hydrocarbonoclastic bacterium Alcanivorax borkumensis. Environ Microbiol 6: 264–273. van Beilen JB, Smits THM, Roos FF, Brunner T, Balada SB, Ro¨thlisberger M, Witholt B (2005) Identification of an amino acid position that determines the substrate range of integral membrane alkane hydroxylases. J Bacteriol 187: 85–91. van Beilen JB, Funhoff EG, Van Loon A, Just A, Kaysser L, Bouza M, Holtackers R, Ro¨thlisberger M, Li Z, Witholt B (2006) Cytochrome P450 alkane hydroxylases of the CYP153 family are common in alkanedegrading eubacteria lacking integral membrane alkane hydroxylases. Appl Environ Microbiol 72: 59–65. Yakimov MM, Timmis KN, Golyshin PN (2007) Obligate oil-degrading marine bacteria. Curr Opin Biotechnol 18: 257–266. Yakimov MM, Golyshin PN, Lang S, Moore ERB, Abraham WR, Lu¨nsdorf H, Timmis KN (1998) Alcanivorax borkumensis gen. nov., sp. nov., a new, hydrocarbon-degrading and surfactant-producing marine bacterium. Int J Syst Bacteriol 48: 339–348.

34 Marinobacter R. Grimaud Institut Pluridisciplinaire de Recherche en Environnement et Mate´riaux, Equipe Environnement et Microbiologie UMR5254 CNRS, IBEAS, Universite´ de Pau et des Pays de l’Adour, Pau, France

1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1290 2 Hydrocarbon Degrading Marinobacters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1290 3 Wax Ester Biosynthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1291 4 Alkane Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1291 5 Strategies for Accessing Alkanes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1292 6 Marinobacter Genome Projects and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1293

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_90, # Springer-Verlag Berlin Heidelberg, 2010

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Marinobacter

Abstract: Bacteria of the genus Marinobacter are omnipresent in the marine environment. Many strains have been recognized as important hydrocarbon degraders in various marine habitats presenting sometimes extreme conditions such as high pH or high salt concentration. Physiological and chemical studies have brought to light the diversity of cellular functions and genetic regulation involved in degradation of aliphatic hydrocarbons by members of the Marinobacter genus. The availability in the near future of genome sequences of five Marinobacter strains will greatly facilitate molecular studies and will offer the possibility of applying ‘‘omics’’ technologies. In due course, these studies should increase our understanding of hydrocarbon degradation in marine environment.

1

Introduction

The genus Marinobacter accommodates Gram-negative, aerobic, motile, nonspore forming, rod-shaped bacteria within the g-subclass of the Proteobacteria. Bacteria of this genus are characterized by the absence of fermentative metabolism, a restricted nutritional profile and the capability of anaerobic respiration on nitrate. The type strain Marinobacter hydrocarbonoclasticus SP17 was isolated for its ability to use alkanes as sole carbon and energy source from chronically oil-contaminated sediment of the French Mediterranean coast (Gauthier et al., 1992). Since then a further 21 Marinobacter species have been described. Marinobacter sp strains were collected in various locations around the world, exclusively from marine or halophilic habitats having diverse physiochemical characteristics including seawater, sediments, marine hot-water springs, sea ice, deep seafloor, saline soil and offshore oil-producing well. Marinobacter species such as M. algicola and M. bryozoorum have been found to be associated with phyto and zooplankton respectively (Romanenko et al., 2005; Green et al., 2006). Consistently with their habitat, most Marinobacter strains are moderate halophiles (See > Chapter 36, Vol. 1, Part 10; > Chapter 54, Vol. 3, Part 4; > Chapter 53, Vol. 3, Part 4).

2

Hydrocarbon Degrading Marinobacters

Hydrocarbon utilization is a common feature in Marinobacter genus. In addition to M. hydrocarbonoclasticus SP17, many strains were found to be alkane degraders, although they have been isolated without using hydrocarbon degradation as a selective pressure. These isolates have been collected from various environments: M. aquaeolei VT8 (subsequently assigned to the specie hydrocarbonoclasticus) was collected from an offshore oil producing well (Huu et al., 1999; Ma´rquez and Ventosa, 2005), M. algicola DG893 from dinoflagellates associated flora (Green et al., 2006), M. maritimus from sea water off the subantarctic Kergelen islands (Shivaji et al., 2005) and M. squalenivorans from oil contaminated marine sediment (Rontani et al., 2003). Marinobacter sp degrading hydrocarbons were also isolated from hydrocarbonoclastic consortia enriched from mangrove sediment or artic sea water (Brito et al., 2006; Deppe et al., 2005). Culture-independent molecular phylogenetic approaches showed that Marinobacters among others specialized hydrocarbonoclastic bacteria belonging to the genera Alcanivorax, Cycloclasticus, Neptunomonas and Thalassolituus were strongly selected in marine sediment amended with hydrocarbons and mineral nutrients (Yakimov et al., 2005).

Marinobacter

34

Some Marinobacter species show remarkable adaptation to hydrocarbon degradation in extreme environments. M. hydrocarbonoclasticus SP17 degrades the solid alkane eicosane at NaCl concentrations up to 3.5 M. Remarkably, eicosane degradation rate and total amount degraded were nearly identical at 0.2 and 2 M (Fernandez-Linares et al., 1996). Alkaliphilic and halophilic hydrocarbon-utilizing Marinobacter species were found in intertidal zone of the coast of the Arabian Gulf in Kuwait. Alkaliphilic isolates degraded hydrocarbons at pH = 11 and halophilic strains at salinity of 3.5% (w/v) (Al-Awadhi et al., 2003). The existence of extremophilic hydrocarbons degraders is important from the bioremediation viewpoint; these organisms could potentially contribute to self-cleaning of oil contaminated alkaline and saline habitats.

3

Wax Ester Biosynthesis

Several Marinobacter species are able to use as sole carbon source isoprenoid compounds such as 6, 10, 14-trimethyl pentadecane-2-one, phytol and squalene which are widely distributed in marine environment. Although the metabolism of isoprenoids by Marinobater sp is mostly aerobic, one strain Marinobacter sp CAB is able to degrade efficiently, 10, 14-trimethyl pentadecane-2-one under both aerobic and denitrifying conditions (Rontani et al., 1997). In N-limited cultures in which ammonium concentrations correspond to that found in marine sediments, aerobic degradation of these compounds leads to the production of relatively large amounts of isoprenoid wax esters. The examination of the ultrastructure of negatively stained cells by electron microscopy showed intracytoplasmic inclusions consisting of accumulated waxes (Rontani et al., 2003). Isoprenoid ester results from the condensation of some acidic and isopronoid alcohol or metabolites produced by their degradation (Rontani et al., 1999). Recently, enzymes of isoprenoid wax esters biosynthesis have been identified and isolated in M. hydrocarbonoclasticus SP17. The activity of two wax ester synthases along with an isoprenoid-specific CoA synthase has been characterized. One of the wax ester synthases exhibits activity toward acyl substrates several orders of magnitude higher than any previously characterized acyl-wax ester synthase (Holtzapple and Schmidt-Dannert, 2007). The ability of Marinobacter sp to use and store widespread marine isoprenoid compounds attest to the well suited physiology of this genus for survival in marine environments. Wax ester accumulation has been also observed in M. hydrocarbonoclasticus SP17 growing on hexadecane. This strain forms a biofilm at alkane-water interface when growing on hexadecane. Cell ultrastructure examination by electron microscopy and lipid analysis revealed that biofilm cells exhibited rounded or irregularly shaped cytoplasmic inclusions and contained large amounts of wax esters. Cells released from the biofilm presented rod shaped inclusions and contained five time less waxes than their sessile counterpart (Klein et al., 2008).

4

Alkane Metabolism

Marinobacters are well represented among the large number of strains isolated from marine environments for their ability to use alkanes as sole carbon and energy sources. Aerobic alkane metabolism was prevailing if not exclusive in these Marinobacters (See > Chapter 3, Vol. 2, Part 2; See > Chapter 23, Vol. 2, Part 5). Studies on the effect of oxygen concentration revealed that

1291

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Marinobacter

heptadecane degradation by Marinobacter nautica strain 617 (subsequently assigned to the Marinobacter genus) is completely abolished below 2.3 mM oxygen (Bonin and Bertrand, 2000). Interestingly, this strain was able to metabolize hydrocarbons under denitrifying and microaerophilic conditions. Nitrate and nitrite were used during growth on heptadecane up to oxygen concentrations corresponding to 30% of air-saturation ( 60 mM) (Bonin et al., 1992). In a pack bed column inoculated with a sample from intertidal mat, alkane crude oil was degraded under microerophilic conditions and Marinobacter sp appeared among the dominant species (Sa´nchez et al., 2006). It seems therefore that the ability to utilize alkane under microaerophlic could be a property shared by many Marinobacter species. Microearophilic and denitrifying condition are very often encountered in marine sediment. The capacity of Marinobacter sp to degrade alkanes under those conditions is evidence of its adaptation for survival and proliferation in hydrocarbon contaminated sediments.

5

Strategies for Accessing Alkanes

Alkanes are nearly insoluble in water and bacterial cells can only grow in aqueous phases. Thus, alkane utilization poses the problem of substrate accessions. Alkane degrading strains have very often been observed sticking at the hydrocarbon water interface and adhesion is thought to be a means to access the hydrocarbon substrate. It is commonly assumed that in order to bind to hydrocarbons bacteria must exhibit a hydrophobic surface. Indeed, M. hydrocarbonoclasticus which adhere to solid and liquid alkanes is highly hydrophobic as attested by a water contact angle of 85 and a surface tension of 25.9 mJ m2 (Bakker et al., 2003). However, adhesion of bacteria to alkanes does not result solely from hydrophobicity and most likely involves multiple interactions. During growth on n-alkanes, fatty alcohols or apolar lipids such as wax esters, triacylglycerol, M. hydrocarbonoclasticus SP17 forms a biofilm at the interface between aqueous medium and the substrates. The concurrent presence of a soluble carbon source (i.e., acetate) does not inhibit biofilm formation on these compounds. In contrast, biofilms were not observed on the non-metabolizable alkanes n-C32, pristane and heptamethylnonane as well as inert substratum such as glass and plastics. The discrimination between metabolizable and non-metabolizable compounds for biofilm formation indicates that at some level, biofilm formation is controlled by the presence of a nutritive interface. In addition, several observations provide evidence that biofilm is required for efficient alkane assimilation and that biofilm formation may be a way to establish and/or promote interfacial access to the alkane (Klein et al., 2008) and unpublished results). The role of surface active compounds (SACs) in alkane degradation by M. hydrocarbonoclasticus SP17 remains unclear as some studies provide inconsistent results. It was found in the extra-cellular medium of a well-agitated culture on the solid alkane eicosane, a high molecular weight SAC possessing an emulsifying activity. Protein and carbohydrate concentration in culture medium are 5 and 10 times higher respectively in culture grown on eicosane than acetate. Proteins and polysaccharides could account for the emulsifying activity detected (Al-Mallah et al., 1990; Fernandez-Linares et al., 1996). Another study demonstrated that there was no emulsification or release of SACs in the bulk medium of a biofilm culture on hexadecane (Klein et al., 2008). Biofilm disruption under strong shaking and release in the medium of biofilm matrix component could account for the emulsifying compounds found in agitated culture. It should be mentioned, however, that

Marinobacter

34

matrix component such as polysaccharides and proteins can be amphiphatic molecules and thus potential emulsifiers, without entailing a role of this property in their physiological functions. Cells of M. hydrocarbonoclasticus SP17 and P. nautica 617 growing on eicosane exhibit blebs at their surface and extracellular vesicles were found in the surrounding medium (Fernandez-Linares et al., 1996; Husain et al., 1997). Similarly, early studies on alkane biodegradation led to the observation of extracellular vesicles during the growth on alkane of A. calcoaceticus 69V (Borneleit et al., 1988). Since then, release of membrane vesicles from the outer membrane have been observed for many gram negative strains. Membrane vesicles derive from the outer membrane and as a consequence possess outer membrane proteins, lipopolysaccharides, phospholipids and periplasm components. However, the relative amount of these components differs in membrane and in vesicles. Membrane vesicles are used for a variety of physiological processes including delivery of toxins to host cells, protein and DNA transfer between bacterial cells, trafficking of cell–cell signals and envelope stress response (Mashburn-Warren and Whiteley, 2006). Schooling and Beveridge showed that membrane vesicles are a common particulate feature of the matrix of Pseudomonas aeruginosa biofilms and a variety of natural multispecies biofilms (Schooling and Beveridge, 2006). The role of membrane vesicles in alkane degradation is still unknown. Their functions could be to expel excess of hydrocarbons as shown for toluene (Kobayashi et al., 2000). Membrane vesicles could be also produced in response to envelope stress caused by contact with the hydrocarbon-water interface (McBroom and Kuehn, 2007).

6

Marinobacter Genome Projects and Research Needs

There are five ongoing genome sequencing projects concerning strains of genus Marinobacter (> Table 1). These strains inhabit extremely diverse marine ecosystems, ranging from deep sea floor to bacterial assemblage associated with algae. The most exotic strain is Marinobacter sp. F02, which was collected from the Juan de Fuca Ridge Middle Valley Vent at a depth of 2400 meters (Edwards et al., 2003). This organism is an obligate Fe-oxidizing autotroph and grows optimally under microaerobic conditions in the presence of an oxygen gradient or anaerobically in the presence of nitrate. Marinobacter sp. ELB17 was isolated from the chemocline waters of a permanently ice-covered lake in the Taylor Valley in Antarctica. Marinobacter algicola DG893 was isolated from the flora associated with the dinoflagellate Gymnodinium catenatum. The symbiotic-like relationship of M. algicola DG893 with its algal host is though to stimulate the dinoflagellate growth and hence the production Toxin Selfish Paralytic (Green et al., 2006). The two other Marinobacter strains being sequenced are M. hydrocarbonoclasticus SP17 and VT8 (formerly aquaeolei) (Huu et al., 1999); (Ma´rquez and Ventosa, 2005). They have been isolated from hydrocarbon contaminated marine environments. It is noteworthy that all sequenced Marinobacter genomes, except the obligate chemolithoautotroph strain, contain genes encoding for putative alkane hydroxylases. Growth on alkanes has been reported for all the tested strains (> Table 1). Whereas the two strains of M. hydrocarbonoclasticus (VT8 and SP17) collected from alkane rich environment possess multiple copies of orthorlogs of alkB and CYP153 genes coding for alkane hydroxylases, M.algicola DG893 and Marinobacter sp ELB17 contain only one copy of alkB homologs and no homologs of CYP153. Alkane assimilation requires much more than simply turning on the corresponding catabolic pathway. Physiological and molecular studies of alkane assimilation carried out on

1293

4.75

4.4

M. Hydrocarbonclasticus Oil well off the coast of Vietnam. VT8. (formerly M. aquaeolei VT8)

Culture of the dinoflagellate Gymnodinium catenatum.

Lake Bonney, ice-covered salted lake in Antarctica.

Deep seafloor, Fe-oxidizing, chemolithoautotrophic

M. algicola DG893

Marinobacter sp. ELB17

Marinobacter s. FO2 2.6

Not available

3.99

M. Hydrocarbonclasticus Contaminated marine sediment of SP17 French Mediterranean coast.

Origin

Genome Size (Mb)

J. Craig Venter Institute

J. Craig Venter Institute

J. Craig Venter Institute

DOE Joint Genome Institute

Genoscope

Sequencing center

Alkane Degradation Genes

Status of the Project

Not tested

Not tested

Sequence not available

In progress

alkaneIn progress monooxygenase

Growth on alkanes 2 alkaneIn progress C16 and C14. monooxygenase

Growth on alkanes 3 alkaneCompleted monooxygenase Automatic 1 CYP 153 annotation Cytochrome

Completed Growth on alkanes 2 alkanemonooxygenase Expert annotation 2 CYP 153 Cytochrome

Degradation of Alkanes

34

Strain

. Table 1 Marinobacter sequencing projects

1294 Marinobacter

Marinobacter

34

different strains established that bacteria interact in a complex way with alkanes. The cellular functions involved are very diverse including production of surface active compounds, interface sensing and adhesion, biofilm formation and precise tuning of intracellular carbon flux. Genome sequence data will permit the application of ‘‘omics’’ technologies which in conjunction with more conventional biochemical and genetic analyses will increase our understanding of the molecular mechanism that underpins alkane degradation in marine environment. The comparison of genomic sequences from different alkanes degrading Marinobacter strains inhabiting diverse marine niches offer the opportunity to unravel the genetic pathway underlying the adaptation to degradation of hydrocarbon in various marine habitats.

References Al-Awadhi H, Al-Hasan RH, Sorkhoh NA, Salamah S, Radwan SS (2003) Establishing oil-degrading biofilms on gravel particles and glass plates. Int Biodeterior Biodegradation 51:181–185. Al-Mallah M, Goutx M, Mille G, Bertrand J-C (1990) Production of emulsifying agents during growth of a marine alteromonas in sea water with eicosane as carbon source, a solid hydrocarbon. Oil Chem Pollut 6:289–305. Bakker DP, Huijs FM, De Vries J, Klijnstra JW, Busscher HJ, Van der Mei HC (2003) Bacterial deposition to fluoridated and non-fluoridated polyurethane coatings with different elastic modulus and surface tension in a parallel plate and a stagnation point flow chamber. Colloids Surf B Biointerfaces 32:179–190. Bonin P, Bertrand JC (2000) Influence of oxygen supply on heptadecane mineralization by pseudomonas nautica. Chemosphere 41:1321–1326. Bonin P, Gilewicz M, Bertrand JC (1992) Effects of oxygen on pseudomonas-nautica growth on n-alkane with or without nitrate. Arch Microbiol 157:538–545. Borneleit P, Hermsdorf T, Claus R, Walther P, Kleber HP (1988) Effect of hexadecane-induced vesiculation on the outer-membrane of acinetobacter-calcoaceticus. J Gen Microbiol 134:1983–1992. Brito EMS, Guyoneaud R, Gon˜i-Urriza M, RanchouPeyruse A, Verbaere A, Crapez MAC, Wasserman JCA, Duran R (2006) Characterization of hydrocarbonoclastic bacterial communities from mangrove sediments in guanabara bay, Brazil. Res Microbiol 157:752–762. Deppe U, Richnow HH, Michaelis W, Antranikian G (2005) Degradation of crude oil by an arctic microbial consortium. Extremophiles 9:461–470. Edwards KJ, Rogers DR, McCollom TM, Wirsen CO (2003) Isolation and characterization of novel psychrophilic, neutrophilic, fe-oxidizing, chemolithoautotrophic a- and g proteobacteria from the deep sea. Appl Environ Microbiol 69:2906–2913.

Fernandez-Linares L, Gauthier M, Acquaviva M, Bertrand JC (1996a) Effect of sodium chloride concentration on growth and degradation of eicosane by the marine halotolerant bacterium marinobacter hydrocarbonoclasticus. Syst Appl Microbiol 19:113–121. Fernandez-Linares L, Acquaviva M, Bertrand JC, Gauthier M (1996b) Effect of sodium chloride concentration on growth and degradation of eicosane by the marine halotolerant bacterium marinobacter hydrocarbonoclasticus. Systematic and Applied Microbiology 19:113–121. Gauthier MJ, Lafay B, Christen R, Fernandez L, Acquaviva M, Bonin P, Bertrand JC (1992) Marinobacter hydrocarbonoclasticus gen. Nov., sp. Nov., a new, extremely halotolerant, hydrocarbon-degrading marine bacterium. Int J Syst Bacteriol 42:568–576. Green DH, Bowman JP, Smith EA, Gutierrez T, Bolch CJS (2006) Marinobacter algicola sp. Nov., isolated from laboratory cultures of paralytic shellfish toxinproducing dinoflagellates. Int J Syst Evol Microbiol 56:523–527. Holtzapple E, Schmidt-Dannert C (2007) Biosynthesis of isoprenoid wax ester in marinobacter hydrocarbonoclasticus dsm 8798: Identification and characterization of isoprenoid coenzyme a synthetase and wax ester synthases. J Bacteriol 189:3804–3812. Husain DR, Goutx M, Bezac C, Gilewicz M, Bertrand JC (1997) Morphological adaptation of pseudomonas nautica strain 617 to growth on eicosane and modes of eicosane uptake. Lett Appl Microbiol 24:55–58. Huu NB, Denner EB, Ha DT, Wanner G, Stan-Lotter H (1999) Marinobacter aquaeolei sp. Nov., a halophilic bacterium isolated from a vietnamese oilproducing well. Int J Syst Bacteriol 49:367–375. Klein B, Grossi V, Bouriat P, Goulas P, Grimaud R (2008) Cytoplasmic wax ester accumulation during biofilmdriven substrate assimilation at the alkane-water interface by marinobacter hydrocarbonoclasticus sp17. Res Microbiol 159:137–144.

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Marinobacter

Kobayashi H, Uematsu K, Hirayama H, Horikoshi K (2000) Novel toluene elimination system in a toluene-tolerant microorganism. J Bacteriol 182: 6451–6455. Ma´rquez MC, Ventosa A (2005) Marinobacter hydrocarbonoclasticus Gauthier et al. 1992 and marinobacter aquaeolei Nguyen et al. 1999 are heterotypic synonyms. Int J Syst Evol Microbiol 55:1349–1351. Mashburn-Warren LM, Whiteley M (2006) Special delivery: Vesicle trafficking in prokaryotes. Mol Microbiol 61:839–846. McBroom AJ, Kuehn MJ (2007) Release of outer membrane vesicles by gram-negative bacteria is a novel envelope stress response. Mol Microbiol 63: 545–558. Romanenko LA, Mikhailov VV, Schumann P, Stackebrandt E, Rohde M, Zhukova NV (2005) Marinobacter bryozoorum sp. Nov. and marinobacter sediminum sp. Nov., novel bacteria from the marine environment. Int J Syst Evol Microbiol 55:143–148. Rontani J-F, Mouzdahir A, Michotey V, Bonin P, Caumette P (2003) Production of a polyunsaturated isoprenoid wax ester during aerobic metabolism of squalene by marinobacter squalenivorans sp. Nov. Appl Environ Microbiol 69:4167–4176. Rontani JF, Bonin PC, Volkman JK (1999) Production of wax esters during aerobic growth of marine bacteria

on isoprenoid compounds. Appl Environ Microbiol 65:221–230. Rontani JF, Gilewicz MJ, Michotey VD, Zheng TL, Bonin PC, Bertrand JC (1997) Aerobic and anaerobic metabolism of 6,10,14-trimethylpentadecan-2-one by a denitrifying bacterium isolated from marine sediments. Appl Environ Microbiol 63:636–643. Sa´nchez O, Ferrera I, Vigue´s N, Oteyza TGd, Grimalt J, Mas J (2006) Role of cyanobacteria in oil biodegradation by microbial mats. Int Biodeterior Biodegradation 58:186–195. Schooling SR, Beveridge TJ (2006) Membrane vesicles: An overlooked component of the matrices of biofilms. J Bacteriol 188:5945–5957. Shivaji S, Gupta P, Chaturvedi P, Suresh K, Delille D (2005) Marinobacter maritimus sp nov., a psychrotolerant strain isolated from sea water off the subantarctic kerguelen islands. Int J Syst Evol Microbiol 55:1453–1456. Yakimov MM, Denaro R, Genovese M, Cappello S, D’Auria G, Chernikova TN, Timmis KN, Golyshin PN, Giluliano L (2005) Natural microbial diversity in superficial sediments of milazzo harbor (sicily) and community successions during microcosm enrichment with various hydrocarbons. Environ Microbiol 7:1426–1441.

35 A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas J. I. Jime´nez,{ . J. Nogales2,{ . J. L. Garcı´a2 . E. Dı´az2,* 1 Department of Microbial Biotechnology, Centro Nacional de Biotecnologı´a-Consejo Superior de Investigaciones Cientı´ficas, Madrid, Spain 2 Department of Molecular Microbiology, Centro de Investigaciones Biolo´gicas-Consejo Superior de Investigaciones Cientı´ficas, Madrid, Spain *[email protected] 1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1298

2 2.1 2.1.1 2.1.2 2.2 2.3 2.4 2.5 2.6 2.7 2.8 2.9

Aromatic Catabolic Pathways in Pseudomonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1299 The b-Ketoadipate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1302 The Protocatechuate Branch . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1302 The Catechol Branch . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1303 The Phenylacetyl-CoA Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1304 The Homogentisate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1305 The Gentisate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1305 The Homoprotocatechuate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1306 The Nicotinate Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1306 The Meta-Cleavage Pathways that Generate 2-Hydroxypentadienoate . . . . . . . . . . . . 1307 The Hydroquinone Pathway . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1308 Other Meta-Cleavage Pathways . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1308

3

Global Cellular Responses to Aromatic Compounds in Pseudomonas . . . . . . . . . 1308

4

Biotechnological Applications of the Metabolism of Aromatic Compounds in Pseudomonas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1315

5

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1318

{

J. I. Jime´nez and J. Nogales attributed equally to this work

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_91, # Springer-Verlag Berlin Heidelberg, 2010

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A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

Abstract: The genetic, and the more recent genomic, proteomic, and metabolomic, approaches that have been undertaken to study the catabolism of aromatic compounds in different Pseudomonas strains have contributed significantly to the acceleration and completion of our understanding on different aspects of the physiology, ecology, biochemistry, and regulatory mechanisms underlying a secondary metabolism that allows the use of this highly abundant carbon source by some bacteria. Comparative genomics suggests that the overall organization of catabolic clusters is conserved across the Pseudomonas genus. However, species-specific and strain-specific variations account for differences in gene arrangements, substrate specificities, and regulatory elements. Moreover, genomic analyses point to the existence of parologous genes likely involved in the degradation of aromatic compounds, suggesting that our current knowledge about the degradative potential of Pseudomonas is still far from complete. On the other hand, many aromatic compounds, e.g., hydrocarbons and phenolic compounds, simultaneously serve as potential nutrients to be metabolized by bacteria but also as cellular stressors. The transcriptomic and proteomic approaches carried out with some Pseudomonas strains provide some light on the biodegradation versus stress dilemma. The increased use of the “omic” techniques, together with the genome-scale metabolic reconstructions developed for some Pseudomonas strains, will certainly contribute significantly to unravel the intricate regulatory and metabolic networks that govern the biodegradation of aromatic compounds, as well as their distribution and ecophysiological relevance. All the basic knowledge generated so far about the metabolism of aromatic compounds in Pseudomonas paves the way for a wealth of biotechnological applications, e.g., bioremediation, biotransformations, biosensors, etc., and it is of great potential in Synthetic Biology. Therefore, Pseudomonas becomes a paradigmatic bacterial genus both for increasing basic knowledge and for applied research within the field of aromatic compounds degradation.

1

Introduction

The g-Proteobacteria of the genus Pseudomonas are widespread colonizers of soil and water, plant leaves and roots, and animals and humans. Thus, P. putida, P. fluorescens, P. stutzeri, and P. mendocina are saprophytic and plant-colonizers with a high potential to degrade organic compounds. P. syringae, P. aeruginosa and P. entomophila are well-known pathogens of plants, humans, and insects respectively, although they can show a free lifestyle too. So far, the genomes of 18 Pseudomonas strains belonging to 7 different species, i.e., P. putida (strains KT2440, GB1, F1 and W619), P. entomophila (strain L48), P. stutzeri (strain A1501), P. fluorescens (strains SBW25, Pf-01 and Pf-5), P. aeruginosa (strains PAO1, PA7, PA14, PACS2, and 2192), P. mendocina (strain ymp), and P. syringae (strains 1448A, B728a, and DC3000), are known. The metabolic versatility and high potential to adapt to different environmental conditions are reflected by the sizes of their genomes (usually >6 Mb), which contain large sets of genes involved in carbon source utilization and adaptation, and explain the ubiquity of this genus (dos Santos et al., 2004, Yan et al., 2008). Among Pseudomonas strains, P. putida KT2440 is probably the best-characterized saprophytic laboratory Pseudomonad that has retained its ability to survive and function in the environment. This bacterium is a plasmid-free derivative of P. putida mt-2, a toluene/xylene degrading strain, that is mainly known or its ability to degrade aromatic compounds and for being an ideal host for expanding the range of substrates that it can degrade and/or biotransform in added-value

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

35

products through the recruitment of genes from other microorganisms. Since P. putida KT2440 is able to colonize the rhizosphere of a variety of crop plants, it is being used also to develop new biopesticides and plant growth promoters (dos Santos et al., 2004). Due to their metabolic versatility, Pseudomonas strains are engaged in important metabolic activities in the environment, particularly in element cycling, and they have often been used as paradigmatic microorganisms to study biodegradation of biogenic and xenobiotic pollutants, such as many aromatic hydrocarbons. In this chapter, we provide a genome-based comparison analysis of the catabolic potential of Pseudomonas species toward aromatic compounds. We then analyze the global response of some Pseudomonas strains when metabolizing aromatic compounds, many of which are major stress factors. Finally, we present some current biotechnological applications regarding the catabolism of aromatics in Pseudomonas.

2

Aromatic Catabolic Pathways in Pseudomonas

The number of aromatic compounds that are mineralized by Pseudomonas strains is overwhelming, covering from biogenic common carbon sources, e.g., benzoate, phenylacetate, aromatic amino acids, etc., to toxic compounds of natural, e.g., toluene, styrene, naphthalene, phenol, etc., or anthropogenic, e.g., polychlorobiphenyls (PCBs), dioxins, nitrotoluenes, etc., origin. The degradation of aromatic compounds constitutes a catabolic funnel where a wide variety of peripheral pathways channel structurally diverse substrates into a few central intermediates (usually dihydroxybenzenes or dihydroxyaromatic acids), which are then ringcleaved and converted to tricarboxylic acid (TCA) cycle intermediates through the corresponding central pathways. In this section, we give an overview of the peripheral and central pathways identified by orthologous comparison analysis of the 18 complete chromosomes of Pseudomonas strains accessible at the NCBI server (http://www.ncbi.nlm.nih.gov/sites/entrez? db¼genomeprj). Our genomic survey revealed, at least, 11 different central pathways to which many different peripheral pathways converge (> Fig. 1). Most these pathways have been only experimentally confirmed in some strains of P. putida, mainly P. putida KT2440 and P. putida F1, and some of them in P. aeruginosa PAO1 (Jime´nez et al., 2004). Among the seven Pseudomonas species analyzed, P. putida appears to be the most versatile in terms of the ability to degrade aromatic compounds, since it contains 9 out of the 11 identified central pathways, which is in agreement with the wide range of niches that this species can colonize (dos Santos et al., 2004). Nevertheless, the ability to degrade aromatic compounds is a strain-specific feature and several pathways present is some strains are lacking in other strains of the same species (> Fig. 1). Regarding the catabolic pathways, those funneling to homogentisate and those that lead to the two branches (catechol and protocatechuate) of the b-ketoadipate pathway are the most widespread in Pseudomonas, probably reflecting that their substrates (aromatic amino acids, phenylpropenoids, and several aromatic acids and phenols) are common carbon sources in the environment, and suggesting that their assembly was an old evolutionary event. Meta-cleavage pathways are highly diverse, whereas ortho-cleavage is restricted to the b-ketoadipate central pathway, and the phenylacetyl-CoA route is the only aerobic hybrid pathway (it does not involve a typical dioxygenase-mediated ring-cleavage) so far identified in Pseudomonas. Nevertheless, some central intermediates, e. g., catechol, can be degraded via both ortho and meta cleavage (> Fig. 1).

1299

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

(Continued)

35

. Figure 1

1300

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

35

Usually, the aromatic catabolic genes are arranged in gene clusters that also contain dedicated regulatory and transport genes, and in some cases, efflux pump-encoding genes. The uptake of the compound inside the cell and the inducible expression of the catabolic genes are important regulatory issues that determine the degradative efficiency of the corresponding pathway. Most of the predicted transporters of aromatic compounds in Pseudomonas belong to the MFS transport family and they can be accompanied by outer membrane porins. Although aromatic compounds can enter the cells by passive diffusion when present at high concentrations, active transport increases the efficiency and rate of substrate acquisition in natural environments where these compounds are present at low concentrations, and they can be involved in the chemotactic response of Pseudomonas toward some aromatics (Parales et al., 2008). On the other hand, the pathway-specific regulation is carried out at the transcriptional level by devoted regulatory proteins (activators and repressors) that are highly diverse in their structure and mechanism of action and that appear to be evolved independently of the catabolic genes. This effector-specific regulation is, in turn, subordinated to a more general control that adjusts the particular transcriptional output to the physiological status of the cell (Cases and de Lorenzo, 2005; Carmona et al., 2008). It should be noted, however, that the catabolic potential in Pseudomonas is much wider than that summarized in > Fig. 1, since many strains whose genomes have not yet been sequenced are endowed with additional pathways for the degradation of relevant aromatic pollutants, e.g., chloroaromatics, nitroaromatics, heterocyclic aromatics, polyaromatic hydrocarbons, etc. (Williams and Sayers, 1994; Wackett, 2003), and some of them are even able to degrade aromatics under anaerobic (denitrifying) conditions (Song and Ward, 2005). Moreover, a significant fraction of the degradation genes are located on catabolic plasmids (and other mobile genetic elements such as transposons), some of which are large transmissible plasmids that have been completely sequenced, e.g., pWW0 and pWW53 for toluene/ xylene degradation, pDTG1, NAH7, and pND6–1 for naphthalene degradation, pADP-1 for atrazine degradation, pCAR1 for carbazole degradation (Miyakoshi et al., 2007). Although these catabolic plasmids play a major role in the distribution of the ability to degrade and utilize aromatic pollutants among Pseudomonas strains and among strains from other bacterial genera, e.g., Burkholderia, Ralstonia, Sphingomonas, etc., (Van der Meer, 2008), the corresponding catabolic genes have not been included in > Fig. 1. Some considerations about these central and peripheral pathways are presented below; however, a deeper biochemical and genetic analysis can be found in a series of excellent review articles (Harayama and Timmis, 1992; Spain, 1995; Harwood and Parales, 1996;

. Figure 1 Aromatic catabolic pathways and their genetic determinants in different Pseudomonas genomes. The different peripheral pathways that converge in the same central intermediate are indicated. The name of the genes coding for the peripheral and central pathways is also shown. The central pathways pca and tod amn cmt indicate convergence routes at a non-aromatic compound, i.e., b-ketoadipate enol-lactone and 2-hydroxypentadienoate, respectively. The abbreviations of the different Pseudomonas species are as follows: ae, P. aeruginosa; fl, P. fluorescens; en, P. entomophila; me, P. mendocina; pu, P. putida; st, P. stutzeri; sy, P. syringae. The names of the strains are specified in those cases where the genes/pathways were identified only in some strains (but not in all available strains) of a particular species.

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A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

Reineke, 1998; Jime´nez et al., 2004) in the Biocatalysis/Biodegradation Database (http:// umbbd.msi.umn.edu/search/), as well as in other chapters of this book.

2.1

The b-Ketoadipate Pathway

The b-ketoadipate pathway is considered one of the most widely distributed sets of genes for the degradation of aromatic compounds in microbes. This central pathway is composed of two ortho-cleavage branches, one for the degradation of protocatechuate (pca genes) and the other for the degradation of catechol (cat genes). In Pseudomonas, these two branches converge at the intermediate b-ketoadipate enol-lactone, which is finally degraded to TCA cycle intermediates by the activities encoded by the genes: pcaD, codes for the lactonase that generates b-ketoadipate; pcaIJ, codes for the CoA-transferase that produces b-ketoadipyl-CoA; and pcaF, encodes the thiolase that generates succinate and acetyl-CoA (Harwood and Parales, 1996; Jime´nez et al., 2004). The genomic analyses reported here show that this pathway is present in all Pseudomonas genomes studied, with the only exception of that from P. mendocina. Although both the cat and pca branches are usually present in most organisms, the cat genes are known to be absent in some members of the rhizobial/agrobacterial groups (a-Proteobacteria) that metabolize catechol and related substrates through a meta-cleavage route (Harwood and Parales, 1996). Interestingly, the cat branch is also missing in the three available genomes of P. syringae, which also do not contain a typical catechol meta-cleavage route (> Fig. 1).

2.1.1

The Protocatechuate Branch

Many abundant aromatic compounds such as quinate (hydroaromatic), p-hydroxybenzoate, and phenylpropenoids (p-coumarate, caffeate, cinnamate, ferulate, etc.) are degraded through the central intermediate protocatechuate. The protocatechuate pathway is widely distributed in bacteria (Harwood and Parales, 1996; Jime´nez et al., 2004), and we have identified the pca genes in all Pseudomonas species sequenced except in P. mendocina (> Fig. 1). The orthocleavage of protocatechuate by the two-component protocatechuate 3,4-dioxygenase (PcaGH) generates a carboxy-cis,cis-muconate that is converted to b-ketoadipate enol-lactone by the action of the PcaC and PcaD enzymes. The pca genes are arranged in a single cluster in P. fluorescens, whereas they are organized in different clusters in other Pseudomonas strains (Jime´nez et al., 2004). The most widespread peripheral pathway that funnels to protocatechuate is that for p-hydroxybenzoate degradation, and it consists of a single flavoprotein monooxygenase activity encoded by pobA (Harwood and Parales, 1996), a gene that is present in all species that harbor the pca pathway (> Fig. 1) but that, with the exception of the P. aeruginosa strains, is not linked to the pca genes. Phenylpropenoids are common carbon sources present in highly-abundant biological polymers like lignin and suberin (Young et al., 2005). Methoxylated phenylpropenoids, such as ferulate, are first converted, through a CoA-dependent nonb-oxidative pathway encoded by fcs and ech genes (also called hca), to vanillin. Vanillin is further converted into protocatechuate by the action of a dehydrogenase and an O-demethylase encoded by the vdh and vanAB genes respectively (Priefert et al., 2001; Jime´nez et al., 2004) (> Fig. 1). The van genes are conserved among all the Pseudomonas strains, except for P. entomophila, P. mendocina, and

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

35

P. fluorescens Pf-01, whereas the fcs-vdh-ech genes are only present in some P. putida strains (KT2440, F1 and W619), and in P. syringae (> Fig. 1). Other phenylpropenoids require previous enzymatic activities before the CoA-dependent activation of the corresponding aromatic acid. Thus, the catabolism of coniferyl alcohol involves its conversion into ferulic acid by an alcohol dehydrogenase (CalA) and an aldehyde dehydrogenase (CalB) (Priefert et al., 2001). calB orthologs were found in the genomes of some strains of P. putida, and P. syringae, whereas a calA ortholog was only identified in P. syringae and the gene is not linked to calB (> Fig. 1). Interestingly, two additional genes, aat, (encoding a putative b-ketothiolase) and acd (encoding a putative acyl-CoA dehydrogenase) cluster with the fcs-vdh-ech genes, and they could be responsible of an alternative CoA-dependent b-oxidative pathway for phenylpropenoids degradation (Priefert et al., 2001). Many other aromatic compounds, such as p-cresols, phthalates, and flavonoids, are also funneled to protocatechuate in different Pseudomonas isolates (Harayama and Timmis, 1992; Wackett, 2003), but the corresponding genes have not been identified in any of the sequenced strains.

2.1.2

The Catechol Branch

Catechol is formed in the degradation of a wide variety of aromatic compounds, e.g., benzoate, benzylamine, tryptophan, aniline, salicylate, anthranilate, and mandelate, some of which are major pollutants, e.g., dibenzothiophene, dibenzofuran, dibenzo-p-dioxin, fluorene, naphthalene, biphenyl, phenol, benzene, toluene, 4-nitrotoluene, and nitrobenzene (Harayama and Timmis, 1992; Williams and Sayer, 1994; Wackett, 2003; Jime´nez et al., 2004). Although the genes involved in the degradation of this broad spectrum of aromatic compounds have been identified in different members of the Pseudomonas genus, only some of these peripheral pathways were shown to be present in the Pseudomonas genomes under study (> Fig. 1). Benzoate is a key intermediate in the catabolism of several aromatic compounds, e.g., mandelate, and its dihydroxylation/decarboxylation to catechol involves the benABCD genes (> Fig. 1). The ben genes have been identified in the Pseudomonas genomes that carry the cat genes for catechol ortho-cleavage, i.e., the catA gene encoding the catechol 1,2-dioxygenase, and further degradation to b-ketoadipate enol-lactone, i.e., the catB and catC genes encoding the cis, cis–muconate cycloisomerase and muconolactone isomerase, respectively. Although the ben and cat genes are located together in the genomes of P. fluorescens, P. aeruginosa, P. stutzeri, and P. entomophila, they are not clustered in most of the P. putida strains (with the only exception of strain W619). Moreover, some Pseudomonas, such as P. fluorescens Pf-01, contain two unlinked ben-cat clusters, which might represent paralogs that function under different environmental conditions (ecoparalogs). On the other hand, the mdl genes found in the genome of P. aeruginosa convert L-mandelate into benzoate (> Fig. 1) (Rosenberg and Hegeman 1971), but they are not linked to the ben genes. Some other common peripheral pathways that funnel into catechol and that are present in the complete Pseudomonas genomes are those for the degradation of tryptophan and salicylate (> Fig. 1). Tryptophan is converted into anthranilate (2-aminobenzoate) through the kynurenine pathway encoded by the kyn genes (Kurnasov et al., 2003), which are present in the genomes of P. fluorescens strain Pf-5 and in all P. aeruginosa strains (> Fig. 1). Anthranilate is then converted into catechol by the anthranilate dioxygenase encoded by the antABC genes (Bundy et al., 1998). Anthranilate is also a common intermediate formed during the aerobic catabolism of some nitroaromatic compounds and N-heterocycles, e.g., carbazole and

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oxoquinoline (Miyakoshi et al., 2007), and it is a precursor for the Pseudomonas quinolone cell-to-cell signal that controls numerous cellular functions and virulence in P. aeruginosa (Farrow and Pesci, 2007). The cat–ant–ben genes in P. aeruginosa are both functionally and physically associated, and therefore, they constitute an example of supraoperonic clustering (catabolic island). In P. fluorescens Pf-5, the kyn and ant genes are also clustered in the chromosome. On the other hand, a decarboxylating flavoprotein monooxygenase (NahG) converts salicylate (2-hydroxybenzoate), an intermediate formed in the catabolism of aromatic hydrocarbons such as naphthalene, into catechol (You et al., 1991), and a nahG ortholog is present in the genome of P. putida GB1 (> Fig. 1). Phenol, a major environmental pollutant, becomes hydroxylated to catechol by the action of a phenol monooxygenase. A cluster of phc orthologs that code for a putative multicomponent diiron phenol hydroxylase (Leahy et al., 2003) is present in the genome of P. putida GB1 (> Fig. 1). Similar gene clusters have been found either in the chromosome or in plasmids, e.g., pVI150, in other Pseudomonas strains (Powlowski and Shingler, 1994; Santos and Sa´Correia, 2007). Other types of phenol monooxygenases, e.g., monocomponent flavoenzymes, present in some Pseudomonas strains (Kallastu et al., 1998) could not be identified in any of the sequenced Pseudomonas under study. The degradation of many chloroaromatic pollutants, e.g., chlorobenzene, PCBs, etc., leads to chlorocatechols as central intermediates. Chlorocatechols are usually channeled to chlorinated TCA cycle intermediates via a modified ortho cleavage pathway (clc genes) that shares the pcaIJ and pcaF genes with the standard b-ketoadipate pathway (Reineke, 1998). Although the clc pathway is present in different Pseudomonas strains, e.g., P. knackmussii B13 (Reineke, 1998), no clc genes were detected in the genomes of the sequenced Pseudomonas strains. Alternative pathways for the degradation of chloroaromatics have been reported in different Pseudomonas strains. Thus, in Pseudomonas sp. strain CBS-3 4-chlorobenzoate becomes dehalogenated via CoA derivatives that generate p-hydroxybenzoate as final product (Dunaway-Mariano and Babbitt, 1994), whereas in P. aeruginosa strain 142 an oxygenolytic ortho-dehalogenation of chlorobenzoates (ohb genes) has been described (Tsoi et al., 1999). However, ortholog genes encoding these alternative pathways are absent in the complete Pseudomonas genomes.

2.2

The Phenylacetyl-CoA Pathway

The aerobic degradation of phenylacetate differs significantly from the classical strategies that involve the activation of aromatic compounds to dihydroxylated intermediates and then a dioxygenolytic ring-cleavage of the latter. The phenylacetate pathway is a hybrid pathway that combines properties from the aerobic and the anaerobic catabolism of aromatic compounds. Thus, the molecule is first activated through the addition of CoA by a phenylacetate-CoA ligase encoded by the paaE gene. Phenylacetyl-CoA then suffers an oxygenation by a heteromultimeric diiron oxygenase (paaGHIJK gene products) to form a 1,2-dihydrodiol intermediate that is not reoxidized to a dihydroxylated aromatic compound, as usual in standard aerobic degradation pathways, but rather subject to enoyl-CoA isomerization/hydration and nonoxygenolitic ring cleavage by the paaBNE gene products. The hydroxyadipyl-CoA compound formed is then oxidized to b-ketoadipyl-CoA, thus converging with the b-ketoadipate central pathway, and then thiolytically split into succinyl-CoA and acetyl-CoA, by the dedicated paaAC and paaE gene products respectively (Olivera et al., 1998; Nogales et al., 2007).

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

35

The paa genes have been identified in all P putida strains, P. entomophila and P. fluorescens Pf-5 (> Fig. 1), and they show a similar gene arrangement in four putative transcriptional units (Jime´nez et al., 2004). Although similar hybrid pathways for the aerobic degradation of benzoate (box genes) and anthranilate (abm genes) have been described in some facultative anaerobes, these pathways have not yet been reported in the Pseudomonas genus (Gescher et al., 2006). The phenylacetyl-CoA pathway constitutes a catabolon through which many other structurally related compounds, such as phenylethylamine, phenylethanol, styrene, tropate, and n-phenylalkanoates with an even number of carbon atoms, are funneled (Luengo et al., 2001). The ped and pea gene clusters encode the phenylethanol dehydrogenase and phenylethylamine dehydrogenase respectively, that convert phenylethanol and phenylethylamine into phenylacetaldehyde in P. putida U (Arias et al., 2008). The pea cluster is present in all the P. putida strains, P. fluorescens Pf-5 and P. entomophila, whereas the ped cluster is lacking in P. putida W619 and P. entomophila (> Fig. 1). On the other hand, these three Pseudomonas species contain fad orthologs for the b-oxidation of n-phenylalkanoates to phenylacetyl-CoA (Olivera et al., 2001) (> Fig. 1).

2.3

The Homogentisate Pathway

The central metabolite homogentisate is formed in the degradation of the aromatic amino acids phenylalanine and tyrosine (Arias-Barrau et al., 2004). The homogentisate central pathway is encoded by the hmg genes responsible for the extradiol-ring cleavage of homogentisate (hmgA), isomerization of maleylacetoacetate (hmgC), and fumarylacetoacetate hydrolysis (hmgB) into fumarate and acetoacetate (Arias-Barrau et al., 2004). The hmg genes can be found in all Pseudomonas species analyzed (> Fig. 1), in agreement with the widespread of aromatic amino acids as carbon sources. Phenylalanine and tyrosine are converted into homogentisate through a peripheral pathway involving: (1) hydroxylation of phenylalanine to tyrosine by the pterin-dependent phenylalanine hydroxylase (phhAB), (2) deamination of tyrosine to 4-hydroxyphenylpyruvate by the tyrosine aminotransferase (tyrB or phhC), and (3) oxygenation and decarboxylation of 4-hydroxyphenylpyruvate into homogentisate by a 4-hydroxyphenylpyruvate dioxygenase (hpd) (Arias-Barrau et al., 2004; Jime´nez et al., 2004). As with the hmg genes, phh/tyr/hpd orthologs are found in all the Pseudomonas genomes (> Fig. 1), but their organization appears to be species-specific. Thus, the hpd gene is clustered with the hmg genes in P. syringae, P. stutzeri, and P. mendocina, whereas it is linked to the phh (tyrB) genes in P. aeruginosa; on the other hand, the hmg/hpd/tyrB/phh genes are dispersed along the genome in P. putida strains. Although homogentisate is a central intermediate in the catabolism of 3-hydroxyphenylacetate in P. putida U (Arias-Barrau et al., 2004), the corresponding mha genes were not detected in the genomes of the sequenced Pseudomonas strains.

2.4

The Gentisate Pathway

Gentisate is the central compound that funnels several peripheral pathways for the catabolism of aromatic acids, such as salicylate and 3-hydroxybenzoate, and some phenol derivatives, e.g., 3,5- or 2,5-xylenol, and m-cresol (Gao et al., 2005). The gentisate central pathway is

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encoded by the gtd genes responsible for the extradiol-ring cleavage of gentisate (gtdA), isomerization of maleylpyruvate (gtdC), and fumarylpyruvate hydrolysis (gtdB) into fumarate and pyruvate (Jime´nez et al., 2004). The gtdABC orthologs have been found in the genomes of P. aeruginosa and P. entomophila (> Fig. 1). Interestingly, an ortholog of the xlnD gene encoding a 3-hydroxybenzoate-6-monooxygenase that converts 3-hydroxybenzoate into gentisate in P. alcaligenes NCIMB 9867 (Gao et al., 2005) was detected in the P. entomophila chromosome linked to the gtd genes (> Fig. 1).

2.5

The Homoprotocatechuate Pathway

The central intermediate homoprotocatechuate is formed during the catabolism of a number of aromatic amines and hydroxylated aromatic acids, e.g., 4-hydroxyphenylacetate. It is degraded through a meta-cleavage pathway that contains a dehydrogenative route yielding finally succinate and pyruvate. The hpa genes that encode the homoprotocatechuate pathway have been described in P. aeruginosa (Cuskey and Olsen, 1988). The genes are arranged in at least four operons: hpaR and hpaA for transcriptional control, hpaBC for the conversion of 4-hydroxyphenylacetate into homoprotocatechuate, and hpaG1G2EDFXHI for the degradation of homoprotocatechuate to TCA cycle intermediates (Jime´nez et al., 2004). Similar hpa genes were identified in P. fluorescens Pf-5, P. entomophila and P. putida GB1 (> Fig. 1), although their arrangement is slightly different from that found in P. aeruginosa. Biogenic amines can be formed by the hydrolysis of arylsulfate esters through the action of the AtsA arylsulfatase. Tyramine and dopamine are catabolized via a tyramine dehydrogenase and a hydroxyphenylacetaldehyde dehydrogenase that produce 4-hydroxyphenylacetate (from tyramine) and homoprotocatechuate (from dopamine). Octopamine, synephrine, and norepinephrine are also used as carbon sources by P. aeruginosa via the homoprotocatechuate pathway, but the genes involved in these peripheral pathways are still unknown (Cuskey and Olsen, 1988).

2.6

The Nicotinate Pathway

Nicotinic acid is a N-heterocyclic aromatic compound that is widely distributed in nature as part of pyridine cofactors (e.g., NAD and NADP) and alkaloids (e.g., nicotine and anabasine), and it is essential (vitamin B3) for many organisms. The complete nicotinate degradation pathway (nic genes) has been recently elucidated in P. putida KT2440, and it involves a nicotinate molybdohydroxylase (nicAB) and 6-hydroxynicotinate monooxygenase (nicC), a peculiar 2,5-dihydroxypyridine extradiol dioxygenase (nicX) and N-formylmaleamate deformylase (nicD), and a set of reactions that convert maleamate into ammonia and fumarate (nicF and nicE) (Jime´nez et al., 2008). In Pseudomonas, the nic genes are arranged in highly conserved clusters that are only present in P. putida strains (> Fig. 1); which is in agreement with the observation that only P. putida strains were reported to use nicotinic acid as sole carbon source (Jime´nez et al., 2008). Some Pseudomonas strains, e.g., Pseudomonas sp. strain S16, are able to degrade nicotine to 2,5-dihydroxypyridine via the pyrrolidine pathway; however only the hsp gene encoding the 6-hydroxy-3-succinoyl-pyridine has been characterized so far (Tang et al., 2008) and it is not present in the sequenced Pseudomonas genomes.

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35

The Meta-Cleavage Pathways that Generate 2-Hydroxypentadienoate

Although benzoate/catechol is usually degraded by the b-ketoadipate pathway, alternative meta-cleavage routes are also present in some Pseudomonas strains. These meta-cleavage pathways are also responsible for the degradation of some alkylcatechols, e.g., 3-methyl- and 3-ethyl-catechol, and protocatechuate derivatives, e.g., 2,3-dihydroxy-p-cumate, that are formed during the aerobic degradation of toxic aromatic hydrocarbons such as toluene, xylenes, ethylbenzene, and p-cymene (Williams and Sayers, 1994). Two Pseudomonas strains, P. putida mt-2 (KT2440 strain containing plasmid pWW0) and P. putida F1, are well-studied aromatic hydrocarbons-degraders that use different biochemical strategies for the activation of the hydrocarbons to the corresponding catecholic intermediates. Thus, P. putida mt-2 contains a pWW0-encoded catabolic pathway (xyl genes) for the oxidation of the alkyl side chain of toluene, m-xylene, and p-xylene (as well as their alcohol derivatives) to the corresponding (methyl)benzoates, and further dioxygenation of the latter to (methyl)catechols (Harayama and Timmis, 1992; Williams and Sayers, 1994). On the contrary, P. putida F1 contains a chromosomally encoded catabolic pathway (tod genes) for the direct dioxygenation of benzene, toluene, and ethylbenzene to catechol, 3-methylcatechol and 3-ethylcatechol respectively (Choi et al., 2003) (> Fig. 1). Both the xyl and tod pathways convert catechol and its alkyl-derivatives into pyruvate and acetyl-CoA via a meta-cleavage route that generates 2-hydroxypentadienoate (HPD) as a common intermediate (Choi et al., 2003). The simultaneous induction in toluene-grown P. putida mt-2 cells of two competing benzoate catabolic pathways, i.e., the chromosomally encoded ben–cat pathway and the pWW0-encoded meta-cleavage pathway, can be explained as the result of cross-regulation between the specific XylS and BenR transcriptional regulators (Domı´nguez-Cuevas et al., 2006). Nevertheless, extended growth of strain mt-2 in benzoate favors the loss of the plasmid pWW0, and therefore, the selection of the ortho-cleavage over the meta-cleavage pathway. Besides benzene, toluene, and ethylbenzene, P. putida F1 is able to degrade p-cymene. The oxidation of the methyl group of p-cymene to produce p-cumate and the ring dioxygenation of the latter to produce 2,3-dihydroxy-p-cumate is encoded by the chromosomally located cym and cmt gene clusters respectively (Eaton, 1997) (> Fig. 1). The further meta-cleavage of 2,3-dihydroxy-p-cumate by the cmtCDE gene products generates HPD, and therefore, converges with the meta-cleavage of alkylcatechols by the todEF gene products (Choi et al., 2003). Further degradation of HPD to pyruvate and acetyl-CoA requires the cmtFGH gene products in the cmt pathway and their corresponding paralogs, todGHI, in the tod pathway. In this sense, the cym and cmt genes are arranged as a single cluster that is located in the vicinity of the tod cluster, thus generating a supraoperonic clustering (catabolic island) for aromatic hydrocarbon degradation in P. putida F1 (Eaton, 1997). Although extradiol dioxygenases usually require the presence of two ortho- or parapositioned hydroxyl-groups in the aromatic ring (Harayama and Timmis, 1992), there are some exceptions. Thus, in the catabolism of some nitroaromatic compounds, such as nitrobencene and 2-nitrobenzoate, two monohydroxylated central intermediates, i.e., 2-aminophenol, and 3-hydroxyanthranilate respectively, are formed. 2-Aminophenol is the substrate of an extradiol dioxygenase (2-Aminophenol 1,6-dioxygenase) that generates 2-aminomuconic semialdehyde, which is then further channeled to HPD. The amn genes for 2-aminophenol catabolism have been characterized in Pseudomonas sp. AP-3 (Takenaka et al., 2000). In P. flurorescens KU-7, 3-hydroxyanthranilate is substrate of the extradiol

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3-hydroxyanthranilate 3,4-dioxygenase in a meta-cleavage pathway (nba genes) that also generates HPD as intermediate (Iwaki et al., 2007). Interestingly, an amn gene cluster has been identified in the genome of P. putida W619 (> Fig. 1).

2.8

The Hydroquinone Pathway

Hydroquinone (1,4-dihydroxybenzene) is a central intermediate formed during the catabolism of acetophenones, e.g., 4-hydroxyacetophenone in P. fluorescens ACB, p-ethylphenol in P. putida JD1, and 4-nitrophenol in P. fluorescens ENV2030 (Moonen et al., 2008a). In P. fluorescens ACB, hydroquinone suffers meta-cleavage by a two-component extradiol hydroquinone 1,2-dioxygenase (hapCD), generating 4-hydroxymuconic semialdehyde (Moonen et al., 2008b), which is then channeled to b-ketoadipate by the action of a semialdehyde dehydrogenase (hapE) and a maleylacetate reductase (hapF). Therefore, the hydroquinone meta-cleavage pathway finally converges with the major ortho-cleavage pathway at the level of b-ketoadipate. The hap gene cluster of P. fluorescens ACB shows similarities with the pnp cluster of P. fluorescens ENV2030 involved in 4-nitrophenol utilization (Moonen et al., 2008a). An ortholog hap gene cluster is found in the genome of P. aeruginosa PA7 (> Fig. 1), although the hapAB genes, responsible for hydroxyacetophenone oxidation to hydroquinone, are not present in this bacterium (Moonen et al., 2008a).

2.9

Other Meta-Cleavage Pathways

The wide diversity of meta-cleavage pathways in Pseudomonas is further extended by the existence of additional meta-cleavage routes. Although the catabolism of protocatechuate via the b-ketoadipate pathway is a widespread feature, protocatechuate can be also subject to meta-cleavage. 4,5-cleavage by the two-component protocatechuate 4,5-dioxygenases (ligAB) appears to be the common way of protocatechuate degradation in Sphingomonas and Comamonas strains. However, there are some examples of this type of extradiol dioxygenases and the corresponding meta-cleavage pathway (oxalomesaconate pathway), in Pseudomonas, e.g., P. straminea NGJ1 (Maruyama et al., 2004). The pro genes of the protocatechuate metacleavage degradation operon in strain NGJ1 have not been found, however, in any of the sequenced Pseudomonas strains. Nevertheless, a galA gene encoding a monocomponent extradiol gallate-dioxygenase has been characterized in P. putida KT2440 (Nogales et al., 2005), and is also present in the genomes of other P. putida strains, suggesting that extradiol cleavage and meta-pathways for the degradation of protocatechuate derivatives such as gallate (3,4,5-trihydroxybenzoate) are also present in the genomes of P. putida strains.

3

Global Cellular Responses to Aromatic Compounds in Pseudomonas

Microorganisms living in environments polluted by aromatic compounds face an interesting paradox. Thus, while many of these compounds can be used as carbon and energy sources, allowing biodegraders to colonize niches refractory to other microbes, they are also stressors for the bacteria above a certain threshold since they are membrane-damaging and

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35

macromolecule-disrupting agents that eventually could lead to cell death (Sikkema et al., 1995). Therefore, when bacteria are exposed to these compounds, they respond by adjusting their gene expression programs to adapt to the new situation. Three types of responses are possible: (1) metabolic programs that include the catabolism of the aromatic compound (if the bacteria can use it as carbon source), and the metabolism of nutrients and intermediates, (2) stress– response programs for adaptation to suboptimal growth conditions, and (3) morphological programs related to shape, transport, and surface chemistry of the cell. All the three major programs are intimately connected and influence each other (Domı´nguez-Cuevas et al., 2006). The recent “omic” approaches allow a system biology view of the metabolic fluxes and gene expression patterns that occur when bacteria face a particular aromatic compound. Most of these studies have been carried out in P. putida. A transcriptomic approach showed the short-term response of P. putida KT2440 (pWW0) citrate-grown cells 15 min after they were challenged with a sublethal concentration of an aromatic hydrocarbon such as toluene (Domı´nguez-Cuevas et al., 2006). Interestingly, only 5% of the reprograming response corresponds to genes involved directly in the metabolism of toluene. The main cellular response involves proteins that are directly or indirectly related to protect the cell against the toxic effects of toluene. These proteins act at three major cellular processes: (1) the cell envelope, (2) the oxidative stress response, and (3) the heat-shock (HS) response. The first defense against environmental stresses is the cell envelope, and accordingly, the strongest response of the cell to a sudden shock of toxic aromatics is observed in functions related to the cell barrier, i.e., lipid metabolism, peptidoglycan synthesis, pili, and flagella, etc., that become repressed (> Fig. 2). That these functions are all inhibited to some degree after the membrane injury may reflect a transient slow-down in motility, chemotaxis, and cell division that consume large amounts of ATP but do not provide traits essential for survival under these conditions, and therefore, repression would be a general strategy to save energy for more useful stress endurance programs (Domı´nguez-Cuevas et al., 2006). Accordingly, a significant number of proteins that contain the conserved GGDEF motif become induced by the toluene-shock. The GGDEF proteins are known to control the cellular levels of c-di-GMP, a cyclic dinucleotide that regulates cell transition to a sessile state after a number of environmental signals (Jenal and Malone, 2006), and their participation in motility toward aromatic (and nonaromatic) compounds in P. putida has been recently shown (Sarand et al., 2008). In contrast to the observed general repression of cell barrier functions, toluene shock in P. putida leads to the induction of genes encoding RND efflux pumps involved in the extrusion of toxic chemicals as a defense mechanism (> Fig. 2). Recently, a new extracytoplasmic function (ECF) sigma factor, RpoT, has been described that controls the expression of envelope-related functions and stress endurance against toluene, including the ttgGHI genes responsible for the main toluene extrusion pump in P. putida DOT-T1E (Duque et al., 2007). The initial damage to the membrane leads to a reduction in electron transport chain activity and an increase in hydrogen peroxide and other reactive oxygen species (ROS) production, which in turn produces a general response to oxidative stress (> Fig. 2). As an immediate consequence, several proteins involved in the response to oxidative damage, e.g., glutathione-related proteins, alkylhydroperoxidases, DNA-metabolism related proteins, are strongly induced (> Fig. 2). On the other hand, oxidative stress is checked by inhibition of iron-acquisition functions, e.g., Fe-pyoverdine complex receptors (FpvA) and other TonB-like receptors, and by slowing down or inhibiting different enzymes involved in metabolic feeding and functioning of the TCA cycle. In contrast, upregulation of the specific metabolic response that leads to toluene catabolism (xyl and ben genes) can be observed (Domı´nguez-Cuevas

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(Continued)

35

. Figure 2

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35

et al., 2006) (> Fig. 2). The synthesis of the new proteome found in cells exposed to toluene could explain the observed upregulation of genes involved in amino acid and nucleotide biosynthesis (> Fig. 2). This hypothesis is supported by the induction of the Tuf-1 elongation factor for protein synthesis and as a molecular chaperone, an inorganic pyrophosphatase (Ppa) as a major provider of energy for the polypeptide-making reactions, and enzymes involved in the synthesis of 6-phosphogluconate (Zwf-1) necessary to replenish some of the

. Figure 2 General scheme of the dual response of P. putida toward a toluene shock. The stress, metabolic or morphological responses are indicated in plain text and italics, respectively. Up-regulated and down-regulated proteins (Domı´nguez-Cuevas et al., 2006) are shown. The abbreviations used are: AceFE (PP_0338–9) Pyruvate dehydrogenase; AhpB, AhpC and AhpF (PP_1084, PP_2439 and PP_2440) Alkylhydroperoxidases; Arg(AFJ) (PP_5185, PP_1079. PP_1346) N-acetylglutamate synthase, ornithine carbamoyltransferase, and glutamate-N-acetyltransferase; AroBK (PP_5078 and PP_5079) 3-dehydroquinate synthase and shikimate kinase; Ben (PP_3161–8) BenABCDKEF proteins for benzoate catabolism; Cco (NOQP) (PP_4250–3) Cytochrome c oxidase cbb3-type subunits; CioA (PP_4651) Ubiquinol oxidase subunit I cyanide insensitive; CspA (PP_0636) Cold shock DNA-binding domain protein; Csu (PP_2358–2363) Type 1 pili subunits; Cyo (ABC) (PP_0812–4) Cytochrome o ubiquinol oxidase subunits; DapB (PP_4725) Dihydrodipicolinate reductase; DnaJ (PP_4726) DnaJ chaperone; DnaK (PP_4727) DnaK chaperone; FadA (PP_2137) 3-ketoacyl-CoA thiolase; FadB (PP_2136) Fatty acid oxidation complex a subunit; Flg (PP_4380, PP_4382, PP_4384–6, PP_4389–91) Flagellar proteins; Fli (PP_4366–7 and PP_4375) Flagellar proteins; FpvA (PP_4217) Outer membrane ferripyoverdine receptor; GabD (PP_0213) Succinatesemialdehyde dehydrogenase; GdhA (PP_0675) Glutamate dehydrogenase; GlcB (PP_0356) Malate synthase; Glt(DB) (PP_5075–6) Glutamate synthase; Gmk (PP_5296) Guanylate kinase; GroEL (PP_1361) Chaperonin 60 kDa; GroES (PP_1360) Chaperonin 10 kDa; GrpE (PP_4728) Heat shock protein GrpE; GshA (PP_0243) Glutamate cysteine ligase; Gst (PP_2023) Glutathione S-transferase; HemN (PP_4264) Oxygen-independent coproporphyrinogen III oxidase; HslVU (PP_5000–1) Heat shock protein HslVU; Hsp20 (PP_3234) Heat shock protein HSP20 family; IbpA (PP_1982) Heat-shock protein IbpA; Lon(1–2) (PP_1443 and PP_2302) ATP-dependent protease La; Mdh (PP_0654) Malate dehydrogenase; Met(EH) (PP_2698 and PP_2375) 5-methyltetrahydropteroyltriglutamate homocysteine S-methyltransferase family protein and 5-methyltetrahydrofolate-homocysteine; Mpl (PP_0547) UDP-N-acetylmuramate:L-alanylgamma-D-glutamyl-meso-diaminopimelate ligase; OmpA (PP_1121) OmpA family protein; OprD (PP_1206) porin D; OprG (PP_0504) outer membrane protein OprG; PbpC (PP_0572) penicillinbinding protein 1C; Pet (ABC) (PP_1317–9) Ubiquinol cytochrome c reductase subunits; Ppa (PP_0538) Inorganic pyrophosphatase; PurU (PP_1943) 3-formyltetrahydrofolate deformylase; PykA (PP_1362) Pyruvate kinase II; RadC (PP_5284) DNA repair protein RadC; Tig (PP_2299) Trigger factor; TonB (PP_3612) putative outer membrane receptor; TtgABC (PP_1386–4) Toluene/ multidrug RND transporter subunits; Ttg2A (PP_0958) Toluene tolerance ABC efflux subunit; Tuf-1 (PP_0440) Translation elongation factor Tu; Usp (PP_2187) Universal stress protein; Uvr (AB) (PP_0483 and PP_1974) Excinuclease ABC A and B subunits; XenA (PP_1254) Xenobiotic reductase A; XenB (PP_1478) Xenobiotic reductase B; Xyl, Xyl proteins for the catabolism of toluene in plasmid pWW0; Zwf-1 (PP_1022) Glucose-6-phosphate 1-dehydrogenase.

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pentose phosphate intermediates needed for the novo nucleotide biosynthesis (Domı´nguezCuevas et al., 2006). Interestingly, Zwf-1 plays a key role in maintaining cellular reducing power, and belongs to the SoxR regulon that controls the oxidative stress response. NADPH is, in fact, the real force that mediates the functioning of many antioxidant enzymes. In this way, it has been shown the upregulation of several enzymes that produce NADPH, e.g., glucose6-phosphate-1-dehydrogenase (Zwf-1), 6-phosphogluconate dehydratase (Edd), glutamate dehydrogenase (GdhA), when cells are cultivated using an aromatic compound as carbon source or under oxidative environmental stress (Zhao et al., 2004; Domı´nguez-Cuevas et al., 2006; Singh et al., 2007). Since some aromatic compounds, such as toluene, disrupt the cytoplasmic membrane releasing lipids and proteins, a HS response leading to the induction of the RpoH regulon can be observed after the toluene insult (Domı´nguez-Cuevas et al., 2006). Thus, chaperones involved in the HS response are up-regulated to cope with the presence of misfolded proteins in the cytoplasm (> Fig. 2). The observed increase in RpoH (s32 factor) in the presence of toluene is due to protein stabilization rather than to gene upregulation, and the triggering of the HS response can then be explained in the context of sigma factor competition. Other HS proteins such as xenobiotic reductase (XenA), cold-shock DNA-binding protein (CspA), and the trigger factor (Tig) were up-regulated, and a few other chaperones were repressed when the cells were exposed to toluene (Domı´nguez-Cuevas et al., 2006) (> Fig. 2). The upregulation of XenA, as well as that of Tuf-1 (see above), agrees with the fact that single mutants deficient in these genes grew more slowly in the presence of toluene and/or were less tolerant to sudden toluene shocks than the parental strain (Segura et al., 2005). The involvement of HS and oxidative stress proteins in response to the toxic effect of aromatic hydrocarbons appears to be a general strategy found also in other aerobic (Agullo´ et al., 2007) and anaerobic (Trautwein et al., 2008) biodegraders. An interesting finding derived from the transcriptomic experiments with P. putida KT2440 (pWW0) confronted with toluene was the observation that the new metabolic and stress programs do not imply significant changes in the levels of the transcriptional machinery (RNA polymerase core or sigma factors) but instead a rapid reassignment of available transcriptional elements from dispensable functions and promoters to functions required for stress endurance. Thus, a certain amount of roaming RNA polymerase available will be assigned to set up these new programs. In this sense, the number of genes that are not expressed in the presence of aromatics may not be specifically repressed, but rather deprived of an otherwise engaged transcriptional apparatus, which is reassigned to express functions that now become compulsory for survival (Dominguez-Cuevas et al., 2006). Nevertheless, this strategy of minimal energy expenditure to adapt to a hostile environment, i.e., aromatic hydrocarbons, may just reflect a short-term response of the cells that initially sense the hydrocarbon as a stressor rather than as a potential nutrient. Alternative mechanisms can prevail in long-term responses to assure survival and metabolic fitness. Thus, it is well known that when E. coli cells face a decrease in the growth potential of the environment, they use a risk-prone foraging behavior that consists of increased motility and the massive induction of transport functions and pathways for the metabolism of unavailable carbon sources (Liu et al., 2005). In a different study, P. putida was shown to sense a mixture of hydrocarbons and other chemicals primarily as an environmental insult and not as a potential nutrient (Vela´zquez et al., 2006). This response was evident from the early transcription of descriptors for the HS response, oxidative damage, DNA injury, and chemical stress. However, the expression of such descriptors decreases over time, and after 180 min the cells adapt to the new conditions following a

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successful reprograming of their proteome and triggering a metabolic program for the degradation of available carbon sources. These results suggest that metabolism of noxious carbon sources has evolved independently of the detoxification mechanisms, and that the genetic networks that make stress responses prevail over metabolic programs have been selected as a clear evolutionary benefit (Vela´zquez et al., 2006). As observed with aromatic hydrocarbons, P. putida exposure to a phenol shock also triggers an important stress program that was monitored by proteomics. The increased formation of ROS induced by phenol will lead to the activation of defense mechanisms against oxidative stress, including upregulation of AhpC, SodB. Tpx, and Dsb proteins. In addition, the upregulation of GrpE, HtpG, DnaK, ClpB, GroEL, UspA, and Tig, all involved in cell protection against a variety of stresses, may confer an increased cell protection and enhanced efficiency of cell repair mechanisms against phenol-induced damage (Santos et al., 2004; Santos et al., 2007). In general, most of the proteins putatively involved in solute transport were down-regulated upon cell exposure to phenol, including the outer membrane OprF porin. A relevant exception is the TolC protein that could function as the outer membrane component of different multidrug efflux systems involved in solvent detoxification (Santos et al., 2004; Santos et al., 2007). The two-component regulatory system ColSR has been shown to regulate phenol tolerance in P. putida, most probably by adjusting permeability of the membrane to phenol (Kivistik et al., 2006). A higher energy demand in phenol-stressed cells to cope with the energetically expensive short-term adaptation mechanisms is consistent with the upregulation of TCA cycle dehydrogenases that generate NADH (Icd, SucA) or FADH2 (SdhA), and enzymes involved in amino acid biosynthesis to restore amino acid pools depleted after the shift to stressful conditions. The upregulation of PckA in response to phenol may indicate that the TCA cycle is directed toward the formation of phosphoenolpyruvate that eventually leads to an increase in peptidoglycan synthesis (Santos et al., 2007). Inhibition of nucleotide biosynthesis (downregulation of PurM, PurL, PyrH) may additionally contribute to the funneling of the available ATP to crucial processes for cell adaptation to phenol stress (Santos et al., 2004). Interestingly, the levels of response to an identical concentration of phenol may differ among two different Pseudomonas strains, e.g., KT2440 and M1 strains, or within the same strain when different alternative carbon sources are used, e.g., succinate and pyruvate, reflecting both differences in the energetic status and fitness of the cells to respond to phenol aggression and differences in the level of stress felt by different cell populations (Santos et al., 2007). Although the maximal induction of adaptation mechanisms is registered during the early response to the chemical insult, stress responses can also be detected in cells already adapted to the new stressing environment. The metabolic and stress programs in P. putida cells growing in glucose in the presence of toluene have been studied by proteomics and transcriptomics (Segura et al., 2005; Volkers et al., 2006; del Castillo and Ramos, 2007). Interestingly, glucose metabolism alleviates the toxic effect of toluene and the extensive reprograming observed when the cells face only toluene (already mentioned) cannot be observed in the presence of toluene plus glucose (del Castillo and Ramos, 2007). The interplay between energy-producing and –consuming processes in P. putida in the presence of glucose and toluene reveals the upregulation of the TCA-cycle enzymes and toluene efflux pumps, and the downregulation of NADP(H) consuming systems (e.g., Fab proteins for fatty acid biosynthesis) as well as that of the proton-consuming ATP synthase. The formation of NADH by the TCA cycle enzymes and the inhibition of the membrane ATP synthase will counteract the dissipation of the proton motive force caused either by the leakage of protons across the membrane after toluene

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accumulation or by the proton-driven extrusion of this hydrocarbon by the cognate efflux pumps. Moreover, P. putida cells counteract toluene stress by preventing its influx through the OprF outer membrane channel (down-regulated) and by increasing membrane stabilization through higher levels of the OprH porin (Volkers et al., 2006). Proteomic studies of Pseudomonas strains when cultured in the presence of different aromatic compounds as sole carbon source reveal also the induction of metabolic and stress programs. On the one hand, cells induce the catabolic pathway involved in the mineralization of the corresponding aromatic compound (enzymes and transport proteins) and modulate the levels of several energy-related enzymes, which will be adapted to funnel the final products of the aromatic catabolic pathways to the central metabolism. As an example of this reprograming, the levels of pyruvate dehydrogenase decreased significantly when P. putida KT2440 is cultured in benzoate as the only carbon source. Since acetyl-CoA and succinate are the final products of benzoate degradation via the b-ketoadipate pathway (see above), it is suggested that the overproduction of acetyl-CoA diminished the role of pyruvate dehydrogenase, which produces acetyl-CoA from pyruvate. The overproduction of acetyl-CoA during phenylethylamine degradation up-regulates two metabolic enzymes, i.e., isocitrate lyase from the glyoxylate shunt and acetoacetyl-CoA thiolase that generates acetoacetyl-CoA and CoA for fatty acid and phospholipid biosynthesis (Kim et al., 2006). Several enzymes involved in protein synthesis were down-regulated in benzoate- and p-hydroxybenzoate-grown P. putida cells, which might reflect a decreased metabolic rate (growth rate) in comparison with that of cells grown in succinate. On the other hand, a number of proteins involved in detoxification, oxidative stress response, and protein folding mechanisms were identified in cells growing in aromatic compounds. In this sense, proteins such as AhpC, GrpE, or DnaK are up-regulated when P. putida KT2440 is cultivated in benzoate or p-hydroxybenzoate, and the GroEL protein is up-regulated in cells growing in phenylalanine (Kim et al., 2006). In Pseudomonas sp. DJ-12 cells, DnaK and GroEL proteins were produced in increasing amounts in the presence of 4-hydroxybenzoate (Park et al., 2001). In P. putida PG150, a KT2440 derivative that harbors the phl genes for phenol catabolism, a proteomic study revealed that growth with phenol, a poor carbon source, induced the OprD porin and several ABC transporters involved in the uptake of additional growth substrates such as amino acids, peptides, polyamines, sugars, and phosphate. Moreover, a set of proteins involved in the HS response, e.g., DnaK and XenB, and in the oxidative stress response, e.g., AhpC, Tsa, Tpx, and DsbA-like, were induced by phenol. Interestingly, a similar global response was observed when phenol was substituted by another poor carbon source such as pyruvate, but not when a preferred carbon source, e.g., succinate, was used. Similarly, the carbon catabolite repression of the phenol-degradation genes in the presence of succinate can also be observed for many of the genes involved in transport, stress response, and nucleotide and amino acid metabolism. Therefore, all these results suggest that a particular gene expression program might be not only due to the carbon source itself but also due to the growth rate of the culture and to the energy state of the bacterial cells (Kurbatov et al., 2006). Many pathways for the degradation of aromatic compounds are located in plasmids that can be horizontally transferred between different bacteria, and therefore, they play a major role in the distribution of the ability to degrade and utilize recalcitrant compounds. Plasmid pCAR1 (199 kb), originally discovered in P. resinovorans CA10, is a self-transmissible plasmid that has been conjugationally transferred into P. putida KT2440 conferring to the later the ability to utilize a N-heterocyclic hydrocarbon, carbazole, as its sole carbon and energy source. To study how the carriage of pCAR1 affects the chromosomal transcriptome, a whole-genome

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

35

oligonucleotide microarray for P. putida KT2440 containing pCAR1 was used. It was revealed that the successful expression of the carbazole catabolic operons on pCAR1 is highly dependent on the regulatory network of the host chromosome. Moreover, various responses were exerted during growth with carbazole, all of them mediated by the alternative sigma factors encoded by the genes algT, rpoH, and rpoS. When the transcriptome of succinate-grown cells of P. putida KT2440 was compared with that of P. putida KT2440 (pCAR1) cells, only subtle changes were observed except for the significant induction of the chromosomal parI gene. The parI gene encodes a putative ParA-like ATPase with a N-terminal Xre-type DNA-binding motif, and it could be involved in active partitioning mechanisms of plasmids and segregation of chromosomes (Miyakoshi et al., 2007).

4

Biotechnological Applications of the Metabolism of Aromatic Compounds in Pseudomonas

Since some aromatic compounds and their derivatives are high value-added products, their synthesis by bacterial biocatalysts, which are cheap and regioselectives, is of great biotechnological interest. Due to the fact that the aromatic compound degradation network of P. putida is well-known and the associated metabolic and stress cellular programs are being unraveled, this bacterium has been extensively used as model biocatalyst (Wackett, 2003). Moreover, in silico genome-scale metabolic reconstructions of P. putida KT2440 (iJN746) and P. aeruginosa PAO1 (iMO1056) have been recently developed (Nogales et al., 2008; Oberhardt et al., 2008; Puchalka et al., 2008). These reconstructions can be used as engineering platforms to explore the potential of these bacteria in bioremediation and biocatalysis. We present here some representative examples of the use of P. putida, and some other Pseudomonas strains, for different biotechnological applications related to the metabolism of aromatic compounds (> Table 1). The broad biocatalytic potential of some Pseudomonas strains toward aromatic compounds that are major environmental pollutants makes them highly suitable for bioremediation purposes (> Table 1). This biocatalytic potential can also be used for the bioproduction of value-added compounds such as N-heterocyclic building blocks, e.g., 6-hydroxynicotinic acid, or flavors, e.g., vanillin (> Table 1). The production of polyhydroxyalkanoates (PHA or bioplastics) is an example of an important biotechnological application of P. putida when cultured in chemically defined media containing different aromatic carbon sources, e.g., phenylalkanoates or toxic aromatic hydrocarbons (Garcı´a et al., 1999; O’Leary et al., 2005). Some recombinant derivatives, e.g., those with disruption or deletion of the fadB and/ or fadA genes involved in the b-oxidation multienzymatic complex, have been engineered as bioplastic overproducers. The physicochemical properties of these new polymers (monomer composition and length) can readily be modified by changing the relative proportion of phenylalkanoate precursor added to the culture broth (Luengo et al., 2003). Since the monomeric units of PHA are enantiomerically pure R-3-hydroxyalkanoic acids, they are potentially interesting starting materials for fine chemical synthesis (synthons) and their production has been enhanced both in vivo and in vitro by expressing a PHA depolymerase (Prieto et al., 2007). P. putida is known as an ideal host for expanding the range of aromatic substrates that it can degrade or biotransform in added-value products, through the recruitment of genes from other microorganisms. For instance, recombinant P. putida KT2440 strains have been used in

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. Table 1 Some biotechnological applications derived from the metabolism of aromatics in Pseudomonas

Strain

Biotechnological Process/Approach

Compound

Reference

Bioremediation P. putida JLR11

2,4,6-Trinitrotoluene (TNT)

Van Dillewijn et al. (2007)

P. putida PCL

Naphthalene

Kuiper et al. (2001)

P. putida UWC3

2,4-Dichlorophenoxyacetate

Dejonghe et al. (2000)

P. knackmussii B13

3- and 4-chlorocatechols, 2-aminophenol

Gaillard et al. (2006)

P. pseudoalcaligenes KF707

Polychlorobiphenyls (PCBs)

Taira et al. (1992)

P. putida F1

Benzene, toluene, phenol

Reardon et al. (2000)

P. aeruginosa RW41

4-Chlorobenzenesulfonate

Blasco et al. (2008)

P. putida DOT-T1E

3-Methylcatechol, p-hydroxybenzoate

Rojas et al. (2004)

P. putida S12

Phenol, p-hydroxybenzoate, cinnamate

Wierckx et al. (2008)

P. putida KT2440

6-Hydroxynicotinate

Jime´nez et al. (2008)

P. putida KTOY02

Benzene and toluene cis-diols, chlorobenzene cis-diols

Ouyang et al. (2007)

Pseudomonas sp. NCIB9816

V cis-dihydrodiols catechols, phenols

Resnick et al. (1996)

P. putida U

From n-phenylalkanoic acids

Luengo et al. (2003)

P. putida CA-3

From styrene and phenylacetate

O’Leary et al. (2005)

P. putida F1 (pVAD, pWW0)

Benzene, toluene, ethylbenzene, xylenes, styrene

Lorenzo et al., 2004

P. putida KTH2 (pESOX3)

Dibenzothiophene desulfurization

Gala´n et al. (2000)

P. fluorescens F113L::1180

Polychlorobiphenyls (PCBs)

Villacieros et al. (2005)

P. putida KT2442::44ES

3-Chlorobenzoate

Klemba et al. (2000)

Bioproduction of aromatics

PHAs production

Pathway expansion

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

35

. Table 1 (Continued) Strain

Biotechnological Process/Approach

Compound

Reference

P. putida KT2440 (pCAR1)

Carbazole

Miyakoshi et al. (2007)

P. knackmussii B13 SN45P-1

Chloro- and methyl-phenols Bioprotection

Erb et al. (1997)

Pseudomonas sp. Y2 PAL-1

Styrene

Alonso et al. (2003)

P. putida PpG7-JAMA21

Naphthalene, 2-methylnaphthalene

Werlen et al. (2004)

P. putida F1G4

Benzene, toluene, ethylbenzene, xylenes, naphthalene.

Phoenix et al. (2003)

P. putida BX5Pu-LUX14

2,4-Dinitrotoluene (2,4-DNT)

De las Heras et al. (2008)

P. putida KT2440ICS

Styrene

Lorenzo et al., 2004

P. putida KT2440ICB

Biphenyl, 4-chlorobiphenyl

Munthali et al. (1996)

P. putida MCR8

3-Methylbenzoate

Ronchel and Ramos (2001)

Biosensors

Biocontainment

oil biodesulfurization, in biodegradation of phenol, styrene, chlorobenzoates, and carbazole (> Table 1). The finding that the ttgGHI genes, encoding the major solvent efflux pump in the solvent-tolerant P. putida DOT-T1E strain, are located on a large self-transmissible plasmid (pGRT1) that confers solvent resistance to toluene-sensitive Pseudomonas strains, such as P. putida KT2440, is of biotechnological interest because the property of solvent tolerance is a phenotype that can be exploited in the bioremediation of heavily polluted sites or in doublephase industrial processes (Rodrı´guez-Herva´ et al., 2007). Solvent-tolerant P. putida strains, e.g., P. putida DOT-T1E or S12 strains, are specially useful as whole-cell biocatalysts in doublephase systems for the production of toxic compounds that partition in the organic phase of a solvent–water mixture. They have been used for the bioproduction of cinnamate, p-hydroxybenzoate, 3-methylcatechol, and phenol (> Table 1). For instance, the efficient production of phenol from glucose in P. putida S12 was achieved by the introduction of the tyrosine-phenol lyase gene (TPL) from Pantoea agllomerans, followed by a combined approach of targeted genetic engineering, random mutagenesis, antimetabolite selection, and high-throughput screening, resulting in the improved P. putida S12TPL3 biocatalyst. By using comparative transcriptomics, nucleotide sequence analysis and targeted gene disruption, it was shown that upregulation of the early shikimate pathway for tyrosine biosynthesis (aro genes), and possibly a decreased carbon flux through the biosynthesis of tryptophan caused by a mutation in the trpE gene, may lead to the enhanced phenol production. In addition, several catabolic routes connected to the tyrosine biosynthetic pathway, e.g., the homogentisate and

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protocatechuate pathways, were up-regulated, suggesting that high intracellular levels of tyrosine (or its intermediates) accumulate in the S12TPL3 strain, and therefore, that TPL activity is the bottleneck in phenol synthesis (Wierckx et al., 2008) The construction of recombinant strains through combination of catabolic modules of different origins may be of interest not only for genetic optimization and design of novel pathways in microorganisms intended for confined or unconfined release, but also for bioprotection of the microbial communities when toxic products arise in catabolic pathways of target pollutants as dead-end metabolites. The degradation of mixtures of chloro- and methyl-aromatics is a well-studied example of the misrouting of pathway intermediates. In this sense, Pseudomonas sp. B13 SN45P-1, which was engineered through rational and spontaneous genetic approaches, was able not only to degrade mixtures of chloro- and methyl-phenols when delivered as shock loads, but also to protect micro-flora and –fauna of the waste treatment plant from lethal effects of the pollutants, thereby assuring the maintenance of the waste treatment process itself (Erb et al., 1997). Small molecule binding is the signal for triggering transcriptional activation for most regulators of biodegradation pathways or solvent pumps, and the regulator/promoter regulatory couples have been utilized in the design of Pseudomonas whole-cell biosensors for a wide range of target compounds, e.g., styrene, naphthalene, benzene, toluene, ethylbenzene, PCBs, and others (> Table 1) (Carmona et al., 2008). A step forward is the selection/design of modified effector-binding sites in mutant regulatory proteins, e.g., XylR, DmpR, XylS, or NahR, to increase or enhance effector recognition specificities (Galvao and de Lorenzo, 2006). As an example, a variant of the toluene-recognizing transcriptional regulator XylR has been obtained that responds to 2,4-dinitrotoluene and it has been used for the production of P. putida strains that emit light upon exposure to residues of explosives in a soil microcosm (de las Heras et al., 2008). Regulatory circuits with predefined effector specificities and delineated DNA-binding abilities will also become essential parts for the rational design of regulatory networks in synthetic biology (Galvao and de Lorenzo, 2006). In this sense, regulators that respond to predetermined chemical species can be paramount for the setting of genetic traps to survey new enzymatic activities in metagenomic libraries (Mohn et al., 2006). On the other hand, biosafety circuits for environmental control have been designed based on aromatic regulatory systems. Various genetic circuits that make recombinant bacteria commit suicide after fulfilling a given function, e.g., biodegradation of an environmental pollutant, or when they escape a predetermined location, e.g., the polluted environment, have been reported (Ronchel and Ramos, 2001), and recombinant P. putida strains capable of degrading biphenyl/chlorobiphenyl or styrene and endowed with an active gene containment system have been described (> Table 1).

5

Research Needs

The genetic, and the more recent genomic, proteomic, metabolomic and fluxomic, approaches that have been undertaken to study the catabolism of aromatic compounds in different Pseudomonas strains have contributed significantly to accelerate and complete our understanding on different aspects of the physiology, ecology, biochemistry, and regulatory mechanisms underlying a secondary metabolism that allows the use of this highly abundant carbon source by some bacteria. Pseudomonas becomes, thus, a paradigmatic bacterial genus both for

A Genomic View of the Catabolism of Aromatic Compounds in Pseudomonas

35

increasing basic knowledge and for applied research in the field of aromatic compounds degradation. Comparative genomics suggest that the overall organization of catabolic clusters is conserved across the Pseudomonas genus. However, species-specific and strain-specific variations account for differences in gene arrangements, substrate specificities, and regulatory elements. A more detailed reconstruction of the evolutionary history of the catabolic clusters will benefit from the continued effort to sequence environmentally relevant strains. The existence of a supraoperonic clustering, or even catabolic islands, in some Pseudomonas strains, as well as the frequent location of mobile genetic elements next to the catabolic genes, facilitates the spreading of these chromosomal catabolic genes among several members of the bacterial community. The role of lateral gene transfer as a major mechanism for the adaptation and evolution of novel catabolic abilities toward aromatic compounds in the ecosystem is reinforced when considering that many degradation clusters are located in transmissible plasmids and transposons. However, there are still many issues regarding the catabolism of aromatic compounds that remain unknown or poorly studied and that, and therefore, require future attention. Thus, there is still a gap of knowledge on the dearomatization activities in facultative anaerobic Pseudomonas and on the links between aerobic and anaerobic pathways in strains that inhabit environments with fluctuating oxygen levels. In this sense, the spread and relevance of aerobic hybrid pathways, such as that described in some bacteria for phenylacetate, benzoate, and anthranilate, should be further explored in Pseudomonas. The biochemistry and genetics of the catabolism of some toxic aromatic hydrocarbons (including heterocyclic and polycyclic aromatic compounds) should also be further investigated. Although the number of complete Pseudomonas genomes available is still limited, genomic analyses point to the existence of many paralogous genes likely involved in the degradation of aromatic compounds; this, in turn, raises the question on the physiological role of this genetic redundancy. Whether such paralogous genes may operate under different ecological conditions (ecoparalogs) in bacteria that inhabit multiple niches, or they are standard paralogs and account for the catabolism of yet unknown aromatic compounds, is still an open question that needs to be answered. Moreover, the existence of many genes that are predicted to encode activities related to the catabolism of aromatic compounds, e.g., oxygenases, transporters, etc., points to the existence of additional catabolic pathways the physiological relevance of which needs to be studied, and suggests that our current knowledge about the degradative potential of Pseudomonas is still far from complete. Although some genome-scale metabolic reconstructions have been recently developed for P. putida and P. aeruginosa, this is still an emerging field and much more effort should be devoted to refine these models and generate new ones for other relevant strains. Integrating the effector-specific regulatory circuits into the global regulatory network of the cell will provide some hints about carbon source preferences and the choice of a particular catabolic pathway over competing pathways that degrade the same substrate, and will allow to better understand and redesign the expression of the catabolic clusters. The combination of metabolic data and regulatory signals in an integrated in silico model that can explain the physiological behavior of the cells when confronted with different environmental signals should be the final target to better predict and control the behavior of the biodegraders. Another important aspect that should be stressed in the near future is the transmembrane trafficking of aromatic substrates and metabolites, especially in those situations that involve syntrophic interactions between two or more different partners for the complete

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mineralization of the aromatic substrates. The interplay between regulatory proteins that recognize aromatic compounds, e.g., sensor histidin-kinases, aromatic transporters, and the chemotactic response, deserves future investigation to fully understand the behavioral responses of Pseudomonas to the presence of aromatic compounds. The role and mechanism of action of some nucleotides, e.g., c-di-GMP, as second messengers that control also motility, and perhaps other adaptative responses to toxic carbon sources, should also be studied in the near future. Many aromatic compounds, e.g., hydrocarbons and phenolic compounds, simultaneously serve as potential nutrients to be metabolized by bacteria but also as stressors, since they are membrane-damaging compounds and their metabolism can also generate stress signals within the cell. The transcriptomic and proteomic approaches carried out with some Pseudomonas strains provide some light on the biodegradation versus stress dilemma. Nevertheless, further research should be carried out to better define the timing and content of the metabolic and stress programs. The characterization of auxiliary genes that are not present in the cognate catabolic clusters but that are also involved, either directly or indirectly, in the catabolic pathways will allow to broad our current view on the constellation of genes/proteins essential for the biodegradation of aromatic compounds in bacteria. The increased use of the “omic” techniques, as well as the systems biology approaches for addressing biological complexity from a holistic perspective, and the application of the network theory to biology, will certainly contribute significantly to unravel the intricate regulatory and metabolic networks that govern the biodegradation of aromatic compounds, as well as their distribution and ecophysiological relevance to carbon flux in the environment. The knowledge of the full range of metabolic capabilities of the microorganisms influencing bioremediation, as well as the monitoring of microbial capacities and their distribution at contaminated sites, will allow to predict how these organisms are likely to respond to changes in environmental conditions that will take place during the course of natural attenuation. Finally, all the knowledge generated so far should accelerate the development of bioremediation technologies, e.g., bioaugmentation. Biotransformation processes based on unprecedented enzymes and pathways with novel metabolic capabilities, as well as the design of novel regulatory circuits and catabolic networks of great biotechnological potential in synthetic biology, are now feasible to approach and they should also be accomplished in the near future.

Acknowledgments Work in our laboratory was supported by grants from the Comisio´n Interministerial de Ciencia y Tecnologı´a (GEN2006-27750-C5-3-E, BIO2006-05957, BFU2006-15214-CO3-01, MMA-PR21/06-039/2006/3-11.2, and CSD2007-00005) and Comunidad Auto´noma de Madrid (P-AMB-259-0505).

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36 Genomics of Methylococcus capsulatus J. C. Murrell Department of Biological Sciences, University of Warwick, Coventry, UK [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1328 2 Genome Sequence of Mc. capsulatus (Bath) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1329 3 Gene Mining of the Mc. capsulatus (Bath) Genome . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1330 4 Postgenomic Studies in Mc. capsulatus (Bath) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1331 5 Genomics of Other Methanotrophs and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . 1332

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_92, # Springer-Verlag Berlin Heidelberg, 2010

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Genomics of Methylococcus capsulatus

Abstract: Methylococcus capsulatus (Bath) is probably the best characterized of the aerobic methane oxidizing bacteria (methanotrophs) and has been the ‘‘workhorse’’ organism for researchers studying the biology of methane oxidation for over 40 years. The genome of Mc. capsulatus (Bath) has recently been sequenced and this provides a metabolic ‘‘blueprint’’ which enables further study on one carbon metabolism and regulation of methane oxidation in this bacterium. Subsequently the genome sequences from other methane oxidizing bacteria have, or are soon to be published and this will provide the necessary background information to carry out comparative genomics and to determine experimentally the molecular basis for obligate methanotrophy in many methane oxidizing bacteria.

1

Introduction

Methylococcus capsulatus (Bath) is an obligate methane oxidizing bacterium (methanotroph) which grows on methane as its sole source of carbon and energy. It can also grow on methanol, the first oxidation product of the methane oxidation pathway. It has a very restricted metabolic lifestyle and cannot grow on multi-carbon compounds. Mc. capsulatus (Bath) is a member of the Gammaproteobacteria, commonly classified as a Type I methanotroph because of its carbon assimilation pathways (primarily assimilating carbon at the oxidation level of formaldehyde using the ribulose monophosphate pathway), intracytoplasmic membrane content (bundles of membranes throughout the cell) and phospholipid fatty acid content (reviewed in Trotsenko and Murrell, 2008). It was first isolated by Roger Whittenbury and colleagues from the Roman baths in the City of Bath, UK, about 40 years ago. Its optimum growth temperature is 45 C which is the temperature of the hot springs entering these Roman baths. This methanotroph is probably the most well studied of this fascinating group of organisms, particularly with respect to its carbon assimilation pathways (See > Chapter 2, Vol. 2, Part 2) and methane monooxygenase (MMO), the first enzyme in the pathway of the biological oxidation of methane, a remarkable enzyme which can oxidize the relatively inert methane molecule to methanol at ambient temperatures and pressures ((See > Chapter 17, Vol. 2, Part 4) Hakemian and Rosenzweig, 2007). There has been considerable interest over the years in the use of methane monooxygenase as a biocatalyst and in bioremediation processes (See > Chapter 17, Vol. 2, Part 4; reviewed in Smith and Dalton, 2004). Mc. capsulatus (Bath) has also attracted attention for its biotechnological potential which started in the 1970s where its use was investigated for the large scale production of single cell protein (SCP). The fall in the price of soya protein in the 1980s resulted in loss of interest in methanotrophs as a source of SCP but more recently interest in production of single-cell protein from methane and methanol has been revived in Norway and Denmark for production of added-value protein products as amino acid-balanced feed for farmed fish and other animals. Mc. capsulatus (Bath) is particularly useful for SCP processes since it can be grown in large scale (10,000 L plus) bioreactors to high cell densities and since it is a moderate thermophile, it is less susceptible to contamination than other methanotrophs. In fact it was the interest in the use of Mc. capsulatus (Bath) for production of SCP that was the driving force behind sequencing of the Mc. capsulatus (Bath) genome by Ward et al. (2004).

Genomics of Methylococcus capsulatus

2

36

Genome Sequence of Mc. capsulatus (Bath)

The first published genome sequence of a methanotroph was that of Mc. capsulatus (Bath) (Ward et al., 2004). The genome is 3.3 Mbp arranged as a single circular molecule with 3, 120 predicted encoding sequences (open reading frames, ORFs) and an average mol G + C content of 63%. There are a considerable number of insertion sequence elements (50) throughout the genome suggesting considerable plasticity in the genome. One of these insertion elements can sometimes be found in the gene cluster encoding soluble methane monooxygenase (sMMO). There are also two putative prophages present. At the time of publication, approximately 1,800 of these ORFs were immediately identifiable as similar to proteins of known function. A summary of the general features of the Mc. capsulatus genome is given in > Table 1 (adapted from Ward et al., 2004). The genome of Mc. capsulatus (Bath) is considerably smaller than that of the facultative methanol utilizer Methylobacterium extorquens AM1 (7 Mb) (Chistoserdova et al., 2003) and much closer in size to the genome of Methylobacillus which is an obligate methylotroph. This small genome size presumably reflects the obligate lifestyle of Mc. capsulatus (Bath) (discussed in Chistoserdova et al., 2005). The availability of a considerable amount of information on the physiology and biochemistry of methanotrophs aided annotation of the Mc. capsulatus (Bath) genome. The presence of one copy of the gene cluster encoding soluble methane monooxygenase and multiple copies of genes encoding particulate methane monooxygenase were confirmed and the large number of genes involved in methanol oxidation (mxa gene clusters) and associated accessory genes were also recognizable, largely due to the work of Lidstrom and colleagues with M. extorquens AM1. As predicted, all of the genes encoding enzymes of the ribulose monophosphate pathway, the primary route of carbon assimilation in Mc. capsulatus (Bath) were also observed, as were genes encoding the serine pathway and the Calvin Cycle which probably only play a minor role in carbon assimilation in Mc. capsulatus (Bath). Interestingly multiple pathways for the metabolism of formaldehyde and three homologs of formate dehydrogenase, involved in the dissimilation of carbon to generate reducing power for the initial oxidation of methane and for biosynthesis were observed. The reasons for the multiplicity of primary one carbon enzymes in Mc. capsulatus (Bath) is not fully understood but it may be a ‘‘fail-safe’’ mechanism for an organism that can only grow on methane or

. Table 1 General characteristics of the genome of Mc. capsulatus (Bath) Size of genome in base pairs

3,304,697

Mol % G + C content of DNA

63.6

Number of predicted coding sequences

3, 120

Number of ribosomal RNA operons

2

Number of transfer RNA genes

46

Predicted polypeptides of known function

1,766

Conserved hypothetical proteins encoded

514

Hypothetical proteins encoded

504

1329

1330

36

Genomics of Methylococcus capsulatus

methanol. The presence of genes encoding both soluble and membrane-bound hydrogenases were confirmed, as were the genes encoding the primary routes of ammonia assimilation (glutamine synthetase, glutamate synthase and alanine dehydrogenase) and those involved in nitrogen fixation. The reasons for the ‘‘obligate nature’’ of Mc. capsulatus (Bath) has been of considerable debate for many years. The genome sequence revealed the presence of genes encoding all TCA cycle enzymes but the absence of 2-oxoglutarate dehydrogenase activity in vitro still suggests that the Krebs Cycle cannot operate in Mc. capsulatus (Bath). Interestingly, a small number of putative transport-related genes for uptake of sugars, carboxylates and amino acids are present in the genome and the function of these needs to be tested experimentally to determine if they play any role in uptake of multi-carbon compounds which might be used in small amounts by Mc. capsulatus (Bath). The issue of the obligate nature of methanotrophs has been discussed in detail (e.g., see Shishkina and Trotsenko, 1982; Wood et al., 2004). The presence of a significant number of putative cytochrome genes again suggests flexibility in electron transport and energy metabolism which can now be verified experimentally. The role of copper ions in the regulation of methane monooxygenase expression and activity has also been a longstanding question in methanotroph biology (reviewed in Hakemian and Rosenzweig, 2007; Murrell et al., 2000). Ward et al. (2004) noted the presence of genes encoding putative copper homeostasis, scavenging and transport systems including a P-type ATPase (the Cop system) and a copper ion efflux system (the Cus complex). The presence of genes encoding putative non-ribosomal peptide synthases (NRPS) which may be involved in copper acquisition by Mc. capsulatus (Bath) and a number of putative two component regulator systems, which are probably involved in environmental regulation of metabolic processes in this bacterium, offers a number of interesting targets for future study. Further details of the genome content of Mc. capsulatus (Bath) can be found in Ward et al. (2004) (see also Kelly et al., 2005) which provides a ‘‘metabolic blueprint’’ and for the first time gives insights into the metabolism of an obligate methanotroph. Fortunately there are a number of genetic tools available to carry out mutagenesis and expression studies with Mc. capsulatus (Bath) and other methanotrophs, in order to verify the genomic potential of this methanotroph (e.g., see Csaki et al., 2001, 2003; Sharpe et al., 2007).

3

Gene Mining of the Mc. capsulatus (Bath) Genome

The availability of the raw genome sequence data for Mc. capsulatus (Bath), even prior to publication of the annotated genome sequence, allowed researchers to search for genes of particular interest to them (gene mining). The first example involved the identification of an ORF encoding a cytochrome P450-like enzyme in the Mc. capsulatus genome. This gene was cloned by PCR and expressed in Escherichia coli. Characterization of the purified enzyme revealed that it was a new class of cytochrome P450 with sterol 14a-demethylase activity, thus playing a role in sterol synthesis in Mc. capsulatus (Bath) (Jackson et al., 2002). Subsequent gene mining work by the same researchers who were studying sterol biosynthesis in prokaryotes (sterol biosynthesis is more generally associated with eukaryotes) revealed the presence of an operon containing essential sterol biosynthesis genes in Mc. capsulatus (Bath). This discovery of squalene monooxygenase and oxidosqualene cyclase genes illustrates the power of bioinformatics in gene mining (Lamb et al., 2007). An ORF (MCA2590) was found upstream of the gene encoding MopE, a surface associated protein of Mc. capsulatus, which is a copper-regulated protein that is probably

Genomics of Methylococcus capsulatus

36

involved in sequestering copper in this methanotroph. Subsequent analysis revealed that MCA2590 encoded a novel di-heme cytochrome c peroxidase that is also regulated by the bioavailability of copper ions (Karlsen et al., 2005). Another example of gene mining is that of analysis of isocitrate dehydrogenase (ICDH) from Mc. capsulatus. The putative icd gene from Mc. capsulatus Bath was cloned by PCR into an E. coli expression vector. Over-expression of ICHD in E.coli facilitated the rapid purification and characterization of this enzyme which was a unique NAD+-dependent homotetrameric enzyme, thus illustrating the utility of a genome sequence for biochemical analysis and metabolic studies (Stokke et al., 2007). Metabolic engineering of methanotrophs can also be considered; for example, expression of a bacterial hemoglobin gene in the obligate methanotroph Methylomonas sp 16a facilitated enhanced expression of astaxanthin, presumably by increasing the activity of oxygen-requiring astaxanthin biosynthesis enzymes (Tao et al., 2007).

4

Postgenomic Studies in Mc. capsulatus (Bath)

In order to make best use experimentally of methanotroph genome sequence information, a number of postgenomic technologies need to be established. One of the most powerful ways of analyzing the expression of proteins under different environmental growth conditions is the use of proteomics. Fortunately, high-resolution two dimensional gel electrophoresis techniques have been developed for Mc. capsulatus (Bath) in order to analyze the proteome. For example, the outer membrane sub-proteome has been analyzed by Berven et al. (2006). Twenty eight unique polypeptides were identified from proteins enriched in outer membranes by mass spectrometry and reference to the Mc. capsulatus genome sequence database. Of these, the function and location of six of these polypeptides were previously known. Development of a number of bioinformatics tools allowed the further identification and putative assignment of function to most of the remaining polypeptides, thus illustrating the importance of bioinformatics in postgenomic studies. Membrane proteins represent a particular challenge for proteomic analysis. In addition to the study above, analysis of membrane proteins from Mc. capsulatus (Bath) has also been achieved using polyvinylidene difluoride (PVDF) membranes in conjunction with matrix-assisted laser desorption/ionization mass spectrometry (MALDI-MS) (Chang et al., 2006). Probably the most comprehensive proteomics study on Mc. capsulatus (Bath) to date is that of Kao et al. (2004) who had independently sequenced the genome of Mc. capsulatus (Bath) to 8X coverage. Quantitative proteomics was carried out on cells which had been grown in the presence of low or high concentrations of copper ions in order to analyze differential expression of proteins. Using a cleavable isotope-coded affinity tag (cICAT) technique, 682 differentially-expressed polypeptides were identified. The expression of pMMO polypeptide or sMMO polypeptides under high and low copper growth conditions respectively was confirmed, but more interestingly, the expression of many of the polypeptides of central carbon assimilation pathways were also identified. Another powerful technique which can now be used to complement the proteomics approach is transcriptomics. This will involve the construction and use of whole genome microarrays for Mc. capsulatus (Bath) to analyze levels of mRNA encoded by genes which are differentially expressed in response to different environmental stimuli (e.g., oxygen, nitrogen source, metal ions). All of these ‘omics technologies will need to be backed up with: (1) carefully defined growth experiments in order to provide suitable cells for analysis, (2) carefully defined genetics experiments to mutate target genes, (3) enzyme analysis (possibly after heterologous expression in E. coli), and (4) suitable bioinformatic analyses.

1331

1332

36 5

Genomics of Methylococcus capsulatus

Genomics of Other Methanotrophs and Research Needs

The first methanotroph genome to be published was that of Mc. capsulatus (Bath). At around the same time, the genome of Methylomonas sp. Strain 16a was sequenced by DuPont. Although the genome sequence is not publically available, a number of interesting publications have been forthcoming concerning metabolic engineering using this Type I methanotroph (e.g., see Sharpe et al., 2007; Tao et al., 2007; and references therein). More recently the genome sequence of the first fully authenticated facultative methanotroph Methylocella silvestris, which grows on methane and also on multi-carbon compounds including acetate and other carboxylates (Dedysh et al., 2005; reviewed in Theisen and Murrell, 2005), has been sequenced. Preliminary screening of the raw sequence information suggests a genome size of around 4.2 Mbp, which is larger than that of Mc. capsulatus (Bath), presumably reflecting the facultative nature of this unusual methanotroph. Around 3,600 of the 4,250 putative protein encoding genes have significant similarity to genes in genomes of other Proteobacteria (Chen and Murrell, unpublished). The completed genome sequence will be fully annotated in the near future, enabling comparative genomics with Methylococcus and other methanotrophs. This comparison will help to unravel some of the mysteries as to why most methanotrophs in culture can only grow on one carbon compounds and again will provide a blueprint from which to test a number of hypotheses experimentally. The genomes of Methylomicrobium album BG8 (another typical Type I methanotroph) and Methylosinus trichosporium OB3b (probably the most well-characterized Type II methanotroph) are currently being sequenced and this will provide further data for comparative genomics across four genera of methanotrophs. Very recently some exciting developments in the field of methanotrophy have been seen with the isolation and characterization of three strains in the phylum Verrucomicrobia which remarkably can grow on methane at very low pH and very high temperature (reviewed in Semrau et al., 2008). The complete genome sequence of isolate V4, initially named ‘‘Methylokorus infernorum’’ (recently renamed Methylacidiphilum infernorum), an isolate from Hell’s Gate, a geothermal region of New Zealand, has just been published. This extremely thermophilic, acidophilic methanotroph has a streamlined genome of 2.3 Mbp, 45.5% G + C content and 2473 protein encoding genes and has allowed the reconstruction in silico of central metabolic pathways and pathways of methane oxidation in this fascinating organism. The availability of genome sequences of several methanotrophs opens up whole new avenues of research in the biology of methane oxidation and while it is important not to ‘‘reinvent the wheel,’’ remembering the wealth of literature on the physiology and biochemistry of methane oxidation and one carbon assimilation pathways in these organisms, these genome ‘‘blueprints’’ will enable valuable comparative genomics. This will hopefully answer a number of longstanding questions such as the ‘‘obligate nature of methanotrophy’’ e.g., why do most methanotrophs grow only on one-carbon compounds? Are there any growth conditions under which apparently ‘‘silent’’ genes of central metabolism, such as those encoding 2-oxoglutarate dehydrogenase, are expressed? Are the genes encoding putative transporters for sugars and amino acids expressed at levels that might enable heterotrophic growth? How are copper ions (and other metals such as iron) sequestered and what are the exact mechanisms by which copper ions regulate methane monooxygenase activity and expression? Improved genetic systems which are continually being developed for methanotrophs will also allow metabolic engineering of certain strains and could for example enable the production of high value products alongside SCP in industrial-scale fermentor cultures of methanotrophs.

Genomics of Methylococcus capsulatus

36

References Berven FS, Karlsen OA, Straume AH, Flikka K, Murrell JC, Fjellbirkeland A, Lillehaug JR, Eidhammer I, Jensen HB (2006) Analysing the outer membrane subproteome of Methylococcus capsulatus (Bath) using proteomics and novel biocomputing tools. Arch Microbiol 184: 362–377. Chang C-Y, Liao H-K, Juo C-G, Chen S-H, Chen Y-J (2006) Improved analysis of membrane protein by PVDF-aided, matrix-assisted laser desorption/ionization mass spectrometry. Ana Chimica Acta 556: 237–246. Chistoserdova L, Chen SW, Lapidus A Lidstrom ME (2003) Methylotrophy in Methylobacterium extorquens AM1 from a genomic point of view. J Bacteriol 185: 2980–2987. Chistoserdova L, Vorholt JA, Lidstrom ME (2005) A genomic view of methane oxidation by aerobic bacteria and anaerobic archaea. Genome Biology 6: 208. Csaki R, Hanczar T, Bodrossy L, Murrell JC, Kovacs, KL (2001) Molecular characterisation of structural genes encoding for a membrane bound hydrogenase in Methylococcus capsulatus (Bath). FEMS Microbiol Lett 205: 203–207. Csaki R, Bodrossy L, Klemm J, Murrell JC, Kovacs KL (2003) Cloning, sequencing and mutational analysis of genes involved in the copper dependent regulation of soluble methane monooxygenase of Methylococcus capsulatus (Bath). Microbiology (UK) 149: 1785–1795. Dedysh SN, Knief C, Dunfield P (2005) Methylocella species are facultatively methanotrophic. J Bacteriol 187: 4665–4667. Hakemian AS, Rosenzweig AC (2007) The biochemistry of methane oxidation. Annu Rev Biochem 76: 223–241. Hou S, et al. (2008) Complete genome sequence of the extremely acidophilic methanotroph isolate V4, Methyloacidiphilum infernorum, a representative of the bacterial phylum Verrucomicrobia. Biology Direct 3: 26–51. Jackson CJ, Lamb DC, Marczylo TH, Warrilow AGS, Manning NJ, Lowe DJ, Kelly DE, Kelly SL (2002) A novel sterol 14a-demethylase/ferredoxin fusion protein (MCCYP51FX) from Methylococcus capsulatus represents a new class of the cytochrome P450 superfamily. J Biol Chem 49: 46959–46965. Kao W-C, Chen Y-R, Yi EC, Lee H, Tian Q, Wu K-M, Tsai S-F, Yu SS-F, Chen YJ, Aebersold R, Chan SI (2004) Quantitative proteomic analysis of metabolic regulation by copper ions in Methylococcus capsulatus (Bath). J Biol Chem 279: 51554–51560. Karlsen OA, Kindingstad L, Angelskar SM, Bruseth LJ, Straume D, Puntervoll P, Fjellbirkeland A, Lillehaug JR, Jensen HB (2005) Identification of a

copper-repressible C-type heme protein of Methylococcus capsulatus (Bath). FEBS J 272: 6324–6335. Kelly DP, Anthony C, Murrell JC (2005) Insights into the obligate methanotroph Methylococcus capsulatus. Trends Microbiol 13: 195–198. Lamb DC, Jackson CJ, Warrilow AGS, Manning NJ, Kelly DE, Kelly SL (2007) Lanosterol biosynthesis in the prokaryote Methylococcus capsulatus: insight into the evolution of sterol biosynthesis. Mol Biol Evol 24: 1714–1721. Murrell JC, McDonald IR, Gilbert B (2000) Regulation of expression of methane monooxygenases by copper ions. Trends Microbiol 8: 221–225. Semrau JD, DiSpirito AA, Murrell JC (2008) Life in the extreme: thermophilic methanotrophy. Trends Microbiol 16: 190–193. Sharpe PL, DiCosimo D, Bosak MD, Knoke K, Tao L, Cheng Q, Ye RW (2007) Use of transposon promoterprobe vectors in the metabolic engineering of the obligate methanotroph Methylomonas sp. Strain 16a for enhanced C40 carotenoid synthesis. Appl Env Microbiol 73: 1721–1728. Shishkina VN, Trotsenko YA (1982) Multiple lesions in obligate methanotrophic bacteria. FEMS Microbiol Lett 13: 237–242. Smith TJ, Dalton H (2004) Biocatalysis by methane monooxygenase and its implications for the petroleum industry. In Petroleum Biotechnology: Developments and Perspectives Studies in Surface Science and Catalysis. R Vazquez-Duhalt and R QinteroRamirez (eds.). Amsterdam: Elsevier, pp. 177–192. Stokke R, Madern D, Fedoy A-E, Karlsen S, Birkeland N-K, Steen IH (2007) Biochemical characterization of isocitrate dehydrogenase from Methylococcus capsulatus reveals a unique NAD+-dependent homotetrameric enzyme. Arch Microbiol 187: 361–370. Tao L, Sedkova N, Yao H, Ye RW, Sharpe PL, Cheng Q (2007) Expression of bacterial hemoglobin genes to improve astaxanthin production in a methanotrophic bacterium Methylomonas sp. Appl Microbiol Biotechnol 74: 625–633. Theisen AR, Murrell JC (2005) Facultative methanotrophs revisited. J Bacteriol 187: 4303–4305. Trotsenko YA, Murrell JC (2008) Metabolic aspects of aerobic obligate methylotrophy. Adv App Microbiol 63: 183–229. Ward N, et al. (2004) Genomic insights into methanotrophy: the complete genome sequence of Methylococcus capsulatus (Bath). PLOS Biology 10: 1616–1628. Wood AP, Aurikko JP, Kelly DP (2004) A challenge for 21st century molecular biology and biochemistry: what are the causes of obligate autotrophy and methanotrophy? FEMS Microbiol Rev 28: 335–352.

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37 Roseobacter A. Buchan1 . J. M. Gonza´lez2 1 Department of Microbiology, University of Tennessee, Knoxville, TN, USA [email protected] 2 Department of Microbiology, University of La Laguna, La Laguna, Tenerife, Spain [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1336 2 Evidence for Hydrocarbon Degradation by Roseobacters in Natural Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1336 3 General Features of Roseobacter Genomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1338 4 Ring-Modifying Pathways and Alkane Hydroxylases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1339 5 Summary and Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1342

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_93, # Springer-Verlag Berlin Heidelberg, 2010

1336

37

Roseobacter

Abstract: Members of the Roseobacter lineage of bacteria are prevalent in diverse marine environments where they carry out critical biogeochemical processes. Recent reports, based primarily on culture-independent studies and reviewed here, provide compelling evidence that members of this abundant lineage are involved in hydrocarbon degradation in natural systems. To determine whether cultured representatives possess similar catabolic capabilities, 24 representative Roseobacter genome sequences were searched for genes sharing homology with those known to be involved in the degradation of hydrocarbons and related compounds. Five distinct pathways for the aerobic degradation of aromatic compounds were identified in the genome collection, as were genes encoding alkane hydroxylases and uncharacterized ringcleaving and -hydroxylating dioxygenases. Taken together, these findings suggest Roseobacters, a group historically overlooked with regard to this physiology, may play important roles in the degradation of hydrocarbons at both naturally occurring and elevated levels in marine environments.

1

Introduction

The Roseobacter clade represents a major marine lineage of biogeochemical importance (reviewed in Buchan et al., 2005; Wagner-Do¨bler, 2006). This large and phenotypically heterogenous group of heterotrophic bacteria falls in the order Rhodobacterales within the a-3 subclass of the Proteobacteria. With very few exceptions, the clade is exclusively marine or hypersaline. The group is well represented across diverse marine habitats with highest abundances typically found in coastal environments or in association with phytoplankton. Yet, members have also been identified in marine systems as disparate as hydrothermal vents and polar sea ice. Molecular-based approaches targeting 16S rRNA genes reveal that the Roseobacter clade comprises upwards of 20% of coastal and 15% of mixed layer ocean bacterioplankton communities (Buchan et al., 2005). Furthermore, representatives of this group stand out as one of the most readily cultivated of the major marine lineages making them ideal model organisms for studies aimed at understanding marine bacterial ecology and physiology. Work in this area is facilitated by recent efforts to uncover the genetic repertoire of environmentally relevant marine microbes that have resulted in the production of almost 40 complete, or nearly complete, Roseobacter genome sequences (Brinkhoff et al., 2008).

2

Evidence for Hydrocarbon Degradation by Roseobacters in Natural Environments

Within the last decade, reports providing evidence for a role for Roseobacters in hydrocarbon degradation in marine systems have surfaced. While still relatively small, this body of literature describes compelling data that group members may play pivotal roles in the metabolism of these compounds in diverse natural environments (> Table 1 and references within). Most of these studies are rooted in molecular-based analyses of 16S rRNA gene sequences and do not rely on traditional culturing approaches, which may explain, in part, why this group has been overlooked with respect to this physiology until recently. Culture-based approaches report mixed success with pure-culture biodegradation studies. For example, Brito et al., isolated a collection of eight Roseobacter strains from mangrove sediments in Brazil by demanding growth on pyrene, naphthalene, or fluoranthrene (Brito et al., 2006). All strains were able to

37

Roseobacter

. Table 1 Representation of Roseobacters in bacterial communities exposed to hydrocarbons

Sample source

Treatment

Roseobacter Approacha representation

Reference

North Sea, surface seawaters Enrichment with crude oil

Clone library

>85%

Brakstad et al. (2005)

Gulf of Alaska, crude oilcontaminated surface sediments

Enrichment with crude oil

Clone library

20%

Chang et al. (2000)

Thames Estuary, UK, surface Enrichment with seawater crude oil

Clone library

15%

Coulon et al. (2007)

Heita Bay, Japan, surface seawater

Enrichment with hexadecane

DGGE

n.d.b

Hara et al. (2003)

Etang de Berre, France, crude oil-contaminated microbial mats

None

Clone library

15%

Hernandez et al. (2006)

Etang de Berre, France, crude oil-contaminated microbial mats

Enrichment with benzothiopene

DGGE

n.d.b

Hernandez et al. (2006)

Thames Estuary, UK, surface Enrichment with seawater decane

Clone library

15%

McKew et al. (2007)

Thames Estuary, UK, surface Enrichment with seawater hexadecane

Clone library

21%

McKew et al. (2007)

Thames Estuary, UK, surface Enrichment with alkane Clone seawater mix library

23%

McKew et al. (2007)

Thames Estuary, UK, surface Enrichment with seawater crude oil

Clone library

3%

McKew et al. (2007)

Sub-Antarctic, surface seawaters

Enrichment with crude oil

Clone library

45%

Prabagaran et al. (2007)

Guanabara Bay, Brazil, surface sediments

Selection on pyrene, naphthalene, and fluoranthrene

Isolation

13%

Brito et al. (2006)

Hiroshima Bay, Japan fuel oil None contaminated tidal flat sediments

Isolation

50%

Katayama et al. (2003)

South China Sea, oilfield sediments

Isolation

n.d.

Ying et al. (2007)

None

n.d. = not determined a Clone library = sequence analysis of PCR amplified 16S rRNA gene sequences from total bacterial community; DGGE = denaturing gradient gene electrophoresis analysis of PCR amplified 16S rRNA gene sequences from total bacterial community; Isolation = cultivation of representative bacteria b Relative representation in 16S rRNA gene amplicon pool not determined, but listed as a ‘‘major’’ band apparent in the DGGE profile that was identified by sequence analysis

1337

1338

37

Roseobacter

transform these polycyclic aromatic hydrocarbons with varying levels of ability (>10–100%) in pure culture. Conversely, five Roseobacter strains isolated from fuel oil contaminated tidal bay sediments were unable to grow on pure alkanes (heptane, octane, decane, undecane, dodecane, or hexadecane) or toluene under the conditions tested (Katayama et al., 2003). Caveats attributed to culture-based approaches are particularly relevant to these types of studies, as the appropriate culture conditions for biodegradation in the lab may not adequately reflect favorable conditions found in situ (e.g., concentrations of compounds, whether compounds are provided singly or in mixtures, whether organisms act alone or in concert, etc.) (Van Hamme et al., 2003). The vast majority of studies referenced in > Table 1 demonstrate that Roseobacter populations respond favorably to hydrocarbon amendment, though general trends concerning preferred substrates or environmental conditions are not yet evident. Brakstad et al., found the relative contribution of Roseobacter sequences in bacterial 16S rRNA gene amplicon pools increased from 21 to 89% following exposure to crude oil for 21 days in a near shore water sample and from Table 2. These strains represent a wide phylogenetic and phenotypic diversity and were isolated from diverse environments (Brinkhoff et al., 2008). The most prevalent pathways and/or genes identified in these genomes are reviewed below. DMS/DMSP metabolisms are discussed in a separate section of this text (See > Chapter 27, Vol. 2, Part 5).

4

Ring-Modifying Pathways and Alkane Hydroxylases

Aerobic bacterial degradation of aromatic hydrocarbons typically proceeds through a limited number of pathways that involve incorporation of molecular oxygen via ringhydroxylating dioxygenases and/or ring-cleaving dioxygenases. Until recently, the paradigm for aerobic aromatic compound degradation involved conversion of a wide array of chemical structures to one of a limited number of di- or trihydroxylated intermediates (e.g., protocatechuate, catechol, gentisate, homoprotocatechuate, homogentisate, hydroxyhydroquinone), followed by oxygen-dependent intra- or extradiol cleavage (Harwood and Parales, 1996). However, recent reports have revealed that CoA activation of the benzene ring followed by non-oxygenolytic ring fission may also be common among aerobic bacteria, including Roseobacters (Moran et al., 2007; Zaar et al., 2004). Regardless of the mechanism of ring cleavage, intermediates from these pathways ultimately feed into the TCA cycle. Thus, these compounds typically serve as primary growth substrates for the organisms that utilize them. Oxygenolytic ring-cleaving pathways: protocatechuate, gentisate, and homoprotocatechuate. The protocatechuate branch of the b-ketoadipate pathway, one of the most broadly distributed pathways for the degradation of aromatic compounds in soil microbes (Harwood and Parales, 1996), is prevalent in Roseobacter genomes; 80% of genomes analyzed possess this pathway (> Table 2). With the exception of genes that may coevolve because they encode subunits of a single enzyme (e.g., pcaHG and pcaIJ) and in strains demonstrating species-level identity (i.e., >99% 16S rRNA gene similarity) there is little gene synteny among the genomes and functionally related genes are distributed across multiple loci (Buchan et al., 2004). A distinguishing feature of this pathway in Roseobacters is the presence of a highly conserved open reading frame (ORF; belonging to the PF05853 superfamily) found immediately adjacent to the pcaHG genes that encode for the ortho-cleaving protocatechuate dioxygenase. Reverse

1339

1340

37

Roseobacter

. Table 2 Prevalence of catabolic pathways for degradation of hydrocarbon and related compounds in representative Roseobacter genome sequences Pathways identifieda, e Strainc

pca

gdo

box

hgd

paa

pAH1

RHD, misc

+

+

+

+

+(2)

+

Dinoroseobacter shibae DFL12*

+

Jannaschia sp. CCS1*

+

Loktanella vestfoldensis SKA53

+

+

Oceanicola batsensis HTCC2597

+

+(2)

+(4)

+(2)

+(2)

d

+

+

+

Oceanicola granulosus HTCC2516 Phaeobacter gallaeciensis BS107

+

+

+

Phaeobacter gallaeciensis 2.10

+

+

+

Rhodobacterales bacterium HTCC2150

+

Rhodobacterales bacterium HTCC2654

+b

Roseobacter denitrificans* OCh 114

+

+

+

+ +

+

+

+(2)

+(2)

+

+

+(2)

+

Roseobacter sp. Azwk-3b

+

Roseobacter sp. CCS2

+

Roseobacter sp. MED193

+

+

+

+

+(2)

Roseobacter sp. SK209–2–6

+

+

+

+(3)

+(3)

Roseovarius nubinhibens ISM

+

+(2)

+

Roseovarius sp. 217

+

+(2)

+(3)

Roseovarius sp. HTCC2601

+

+(2)

+(2)

+(2)

+(3)

+(2)

Roseovarius sp. TM1035

+

Sagittula stellata E-37

+

+

+

+

+

+(3)

+(7)

Silicibacter pomeroyi DSS-3*

+d

+(2)d

+

+

+

+(2)

+(4)d

Silicibacter sp. TM1040*

+

+d

+

+

+d

Sulfitobacter sp. EE-36

+

+

+(2)

Sulfitobacter sp. NAS-14.1

+

+

+(2)

0.54

0.88

Frequency

0.79

0.17

0.17

0.42

0.75

Pathway/gene abbreviations are as follows: pca = ortho cleavage of protocatechuate (exception: b meta cleavage of protocatechuate); gdo = cleavage of gentisate; box = benzoyl-CoA pathway; hgd = meta cleavage of homoprotocatechuate; paa = phenylacetyl-CoA pathway; pAH1 = alkane monooxygenase; RHD = ringhydroxylating dioxygenases; misc. = uncharacterized ring-cleaving dioxygenases. Positive signs indicate the presence of ORFs with significant similarity (tBLASTx scores >30) to complete sets of characterized genes for pathways (pca, gdo, box, hgd, paa) or individual genes (pAH1, RHD, misc.) per references provided in the text. Prevalence of multiple pathways/genes is provided in parenthesis; ORFs putatively encoding subunits of a single RHD are only counted once c Refer to Brinkhoff et al. (2008) for information on strain origin *Completed genomes are indicated with an asterisk d Denotes plasmid-encoded genes. Two of the four reports for S. pomeroyi in the RHD, misc. category are plasmidborne e Prevalence of ring-cleaving pathways for Jannaschia sp. CCS1, S. pomeroyi, and Silicibacter sp. TM1040 were previously reported in Moran et al. (2007) a

Roseobacter

37

transcription-PCR analysis of RNA from one isolate, Silicibacter pomeroyi, provides evidence that this ORF is co-expressed with upstream pca genes (Buchan et al., 2004). The absence of this ORF in similar bacterial pca gene clusters from diverse microbes suggests a niche-specific role for its protein product in Roseobacter group members. Three additional oxygenolytic ring-cleaving pathways are found in Roseobacter genomes. Genes encoding proteins of the cleavage pathway for gentisate (Adams et al., 2006) are found in 4 (17%) of the genomes. Interestingly, Silicibacter pomeroyi appears to possess two complete copies of this pathway; one plasmid encoded, the other located on the chromosome (Moran et al., 2004). Characterization of the plasmid-encoded gentisate 1,2-dioxygenase from S. pomeroyi revealed the active protein exists in an unusual homotrimeric conformation (Liu et al., 2007). The meta cleavage pathway of homoprotocatechuate (Roper et al., 1993) is found in 10 (42%) of the genomes analyzed. One single genome sequence, from strain HTCC2654, revealed the presence of genes for the meta cleavage pathway of protocatechuate (Noda et al., 1990) (> Table 2). Genes encoding enzymes for either the ortho or meta cleavage of the widespread intermediate catechol are absent from the Roseobacter genomes. Non-oxygenolytic ring fission pathways: benzoate and phenylacetate. Two ring-cleaving pathways that involve CoA thioesterification of the aromatic ring are found in Roseobacter genomes. The recently elucidated benzoyl-CoA pathway (Zaar et al., 2004) is found in 4 (17%) of the genomes, while the phenylacetic acid pathway (Ferra´ndez et al., 1998) is found in just over half (54%) of the genomes (> Table 2). Ring-hydroxylating and miscellaneous ring-cleaving dioxygenases. Ring-hydroxylating dioxygenases are responsible for preparing a variety of structurally diverse aromatic compounds for ring cleavage (Butler and Mason, 1997). While none of the well-characterized ringhydroxylating dioxygenases (RHD) (e.g., those involved in the degradation of naphthalene, phenanthrene) were identified in the 24 Roseobacter genomes several uncharacterized genes with signature sequences characteristic of RHD were (e.g., PF00355, PF00848, PF00866). Seventeen (71%) of the Roseobacter genomes possess putative ring-hydroxylating dioxygenases. Furthermore, additional putative ring-cleaving dioxygenases (e.g., PF02900, PF04444, PF00755, PF00903) were also present in the genomes and may indicate the presence of novel or poorly studied pathways for aromatic compound degradation. Alkane hydroxylases. Putative medium-chain-length integral membrane alkane hydroxylase genes (similar to pAH1; van Beilan et al., 2007) were found in all but three of the Roseobacter genomes (> Table 2). With the exception of Sagittula stellata, Loktanella vestfoldensis, Roseobacter sp. MED193, Roseovarius sp. HTCC2601, and Roseovarius sp. TM1035, these 21 genomes appear to lack genes for either rubredoxin or rubredoxin reductase, both of which are required electron transfer proteins in characterized alkane hydroxlyase systems from g-proteobacteria and Gram positive bacteria (reviewed in van Beilan et al., 2007). Presumably, alternative electron transfer partners would need to be identified in Roseobacters. Synopsis of genome analyses. Roseobacters harbor a wealth of catabolic pathways for the aerobic degradation of phenolic compounds, a feature that is more reminiscent of soil borne (e.g., Pseudomonas spp., Rhodococcus spp.) rather than marine microbes. The diversity and prevalence of ring-cleaving pathways is impressive in the group as a whole and in several strains, in particular. Sagittula stellata, Jannaschia sp. CCS1, Silicibacter pomeroyi, and strain HTCC2654 possess pathways for the degradation of protocatechuate, gentisate, homoprotocatechuate, benzoate, and phenylacetate. Furthermore, genes showing homology to alkane hydroxylases and ring-hydroxylating and/or additional ring-cleaving dioxygenases are present

1341

1342

37

Roseobacter

in these four strains isolated from geographically and physiochemically diverse environments (Brinkhoff et al., 2008) suggesting these may be exceptionally appropriate for future study.

5

Summary and Research Needs

Several lines of compelling evidence indicate a role for Roseobacters in the aerobic degradation of hydrocarbons. Culture-independent studies suggest Roseobacters in diverse marine systems can be stimulated by enrichment with a range of hydrocarbon-based compounds. The extent of the response and the specific group members or subgroups within the lineage that respond likely depends upon many factors, including composition of the original community, the type and concentration of compound(s) added, and physiochemical nature of the system. Pathways for the oxygen-dependent degradation of aromatic compounds and genes encoding for enzymes involved in alkane degradation are prevalent in Roseobacter genomes, though there is significant diversity in the set of pathways present in a given strain, even those within the same species (e.g., P. gallaeciensis). Given our current limited understanding of the abilities of specific group members to degrade these structurally diverse compounds, generalizations regarding the catabolic capabilities of cultured representatives are not yet feasible. However, the lack of extradiol cleavage pathways for the degradation of hydrocarbons and related compounds may suggest these organisms employ non-traditional approaches for the utilization of these substrates. Research directives aimed at uncovering the genetics and biochemistry of hydrocarbon and related compound catabolism in Roseobacters, particularly those for which genome sequences are available, are needed, as are field studies that incorporate quantitative measurements of specific bacterial groups and employ labeled substrates (tracers). Efforts on both complementary fronts will undoubtedly provide a more conclusive understanding of the contribution of this naturally abundant and physiologically versatile group of microbes to hydrocarbon degradation in marine systems.

References Adams MA, Singh VK, Keller BO, Jia Z (2006) Structural and biochemical characterization of gentisate 1,2dioxygenase from Escherichia coli O157:H7. Mol Microbiol 61: 1469–1484. Biers EJ, Wang K, Pennington C, Belas R, Chen F, Moran MA (2008) Occurrence and expression of gene transfer agent genes in marine bacterioplankton. Appl Environ Microbiol 74: 2933–2939. Brakstad OG, Lødeng AGG (2005) Microbial diversity during biodegradation of crude oil in seawater from the North Sea. Microb Ecol 49: 94–103. Brinkhoff T, Giebel H-A, Simon M (2008) Diversity, ecology, and genomics of the Roseobacter clade: a short overview. Arch Microbiol 189: 531–539. Brito EMS, Guyoneaud R, Gon˜i-Urriza M, Ranchou-Peyruse A, Verbaere A, Crapez MAC, Wasserman JCA, Duran R (2006) Characterization

of hydrocarbonoclastic bacterial communities from mangrove sediments in Guanabara Bay, Brazil. Res Microbiol 157: 752–762. Buchan A, Gonza´lez JM, Moran MA (2005) Overview of the marine Roseobacter lineage. Appl Environ Microbiol 71: 5665–5677. Buchan A, Neidle EL, Moran MA (2004) Diverse organization of genes of the b-ketoadipate pathway in members of the marine Roseobacter lineage. Appl Environ Microbiol 70: 1658–1668. Butler CS, Mason JR (1997) Structure-function analysis of the bacterial aromatic ring-hydroxylating dioxygenases. Adv Microb Physiol 38: 47–84. Chang Y-J, Stephen JR, Richter AP, Venosa AD, Bru¨ggemann J, Macnaughton SJ, Kowalchuk GA, Haines JR, Kline E, White DC (2000) Phylogenetic analysis of aerobic freshwater and marine enrichment cultures efficient in hydrocarbon degradation:

Roseobacter effect of profiling method. J Microbiol Methods 40: 19–31. Coulon F, McKew BA, Osborn AM, McGenity TJ, Timmis KN (2007) Effects of temperature and biostimulation on oil-degrading microbial communities in temperate estuarine waters. Environ Microbiol 9: 177–186. Ferra´ndez A, Min˜ambres B, Garcı´a B, Olivera ER, Luengo JM, Garcı´a JL, Dı´az E (1998) Catabolism of phenylacetic acid in Escherichia coli. J Biol Chem 273: 25974–25986. Grayston SJ, Griffith GS, Mawdsley JL, Campbell CD, Bardgett RD (2001) Accounting for variability in soil microbial communities of temperate upland grassland ecosystems. Soil Biol Biochem 33: 533–551. Hara A, Syutsubo K, Harayama S (2003) Alcanivorax which prevails in oil-contaminated seawater exhibits broad substrate specificity for alkane degradation. Environ Microbiol 5: 746–753. Harwood CS, Parales RE (1996) The b-ketoadipate pathway and the biology of self-identity. Ann Rev Microbiol 50: 553–590. Hernandez-Raquet G, Budzinski H, Caumette P, Dabert P, Le Me´nach K (2006) Molecular diversity studies of bacterial communities of oil polluted microbial mats from the Etang de Berre (France). FEMS Microbiol Ecol 58: 550–562. Katayama Y, Oura T, Iizuka M, Orita I, Cho KJ, Chung IY, Okada M (2003) Effects of spilled oil on microbial communities in a tidal flat. Mar Poll Bull 47: 85–90. Liu D, Zhu T, Fan L, Quan J, Guo H, Ni J (2007) Identification of a novel gentisate 1,2-dioxygenase from Silicibacter pomeroyi. Biotech Lett 29: 1529–1535. McKew BA, Coulon F, Osborn AM, Timmis KN, McGenity TJ (2007) Determining the identity and roles of oil-metabolizing marine bacteria from the Thames estuary, UK. Environ Microbiol 9: 165–176. Moran MA, Belas R, Schell MA, Gonza´lez JM, Sun F, Sun S, Binder BJ, Edmonds J, Ye W, Orcutt B, Howard EC, Meile C, Palefsky W, Goesmann A, Ren Q, Paulsen I, Ulrich LE, Thompson LS, Saunders E, Buchan A (2007) Ecological genomics of marine Roseobacters. Appl Environ Microbiol 73: 4559–4569. Moran MA, Buchan A, Gonza´lez JM, Heidelberg JF, Whitman WB, Kiene RP, Henriksen JR, King GM, Belas R, Fuqua C, Brinkac L, Lewis M, Johri S, Weaver B, Pai G, Eisen JA, Rahe E, Sheldon WM, Ye W, Miller TR, Carlton J, Rasko DA, Paulsen IT, Ren Q, Daugherty SC, Deboy RT, Dodson RJ, Durkin AS, Madupu R, Nelson WC, Sullivan SA, Rosovitz MJ, Haft DH, Selengut J, Ward N (2004)

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Genome sequence of Silicibacter pomeroyi reveals adaptations to the marine environment. Nature 432: 910–913. Noda Y, Nishikawa S, Shiozuka K, Kadokura H, Nakajima H, Yoda K, Katayama Y, Morohoshi N, Haraguichi T, Yamasaki M (1990) Molecular cloning of the protocatechuate 4,5-dioxygenase genes of Pseudomonas paucimobilis. J Bacteriol 172: 2704–2709. Norris TB, Wraith JM, Castenholz RW, McDermott TR (2002) Soil microbial community structure across a thermal gradient following a geothermal heating event. Appl Environ Microbiol 68: 6300–6309. Prabagaran SR, Manorama R, Delille D, Shivaji S (2007) Predominance of Roseobacter, Sulfitobacter, Glaciecola and Psychrobacter in seawater collected off Ushuaia, Argentina, Sub-Antarctica. FEMS Microbiol Ecol 59: 342–355. Pradella S, Allgaier M, Hoch C, Pa¨uker O, Stackebrandt E, Wagner-Do¨bler I (2004) Genome organization and localization of the pufLM genes of the photosynthesis reaction center in phylogenetically diverse marine Alphaproteobacteria. Appl Environ Microbiol 70: 3360–3369. Roper DI, Fawcett T, Cooper RA (1993) The Escherichia coli C homoprotocatechuate degradative operon: hpc gene order, direction of transcription and control of expression. Mol Gen Genet 237: 241–250. Swingley WD, Sadekar S, Mastrian SD, Matthies HJ, Hao J, Ramos H, Acharya CR, Conrad AL, Taylor HL, Dejesa LC, Shah MK, O’Huallachain ME, Lince MT, Blankenship RE, Beatty JT, Touchman JW (2007) The complete genome sequence of Roseobacter denitrificans reveals a mixotrophic rather than photosynthetic metabolism. J Bacteriol 189: 683–690. van Beilan JB, Funhoff EG (2007) Alkane hydroxylases involved in microbial alkane degradation. App Microbiol Biotech 74: 13–21. Van Hamme JD, Singh A, Ward OP (2003) Recent advances in petroleum microbiology. Microbiol Mol Biol Rev 67: 503–549. Wagner-Do¨bler I, Biebl H (2006) Environmental biology of the marine Roseobacter lineage. Ann Rev Microbiol 60: 255–280. Ying J-Y, Wang B-J, Xin D, Yang S-S, Liu S-J, Liu Z-P (2007) Wenxina marina gen. nov., sp. nov., a novel member of the Roseobacter clade isolated from oilfield sediments of the South China Sea. Int J Syst Evol Microbiol 57: 1711–1716. Zaar A, Gescher J, Eisenreich W, Bacher A, Fuchs G (2004) New enzymes involved in aerobic benzoate metabolism in Azoarcus evansii. Mol Microbiol 54: 223–238.

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38 Rhodococcus: Genetics and Functional Genomics M. J. Larkin* . L. A. Kulakov . C. C. R. Allen School of Biological Sciences and The QUESTOR Centre, The Queen’s University of Belfast, Belfast, Northern Ireland, UK *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1346 2 The Genetic Basis of Biodegradation Capability . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1346 3 Genome Size and Composition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1347 4 Discovery of New Catabolic Pathways from the Genome Sequence . . . . . . . . . . . . . . . 1348 5 The Role of Catabolic Plasmids . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1348 6 Catabolic Gene Regulation and Expression . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1349 7 Gene Expression in Response to Environmental Stresses . . . . . . . . . . . . . . . . . . . . . . . . . . . 1350 8 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1351

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_94, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Members of the genus Rhodococcus (See also > Chapter 17, Vol. 3, Part 1) are a very diverse group of bacteria that possess the ability to degrade a large number of organic compounds including some of the most difficult compounds with regard to recalcitrance and toxicity. This is based upon the acquisition of a wide and diverse range of catabolic genes housed in a robust cellular environment. They have very large genomes (up to 9.7 Mbps) and multiple catabolic enzymes. They also harbor many large linear plasmids that contribute to their substrate diversity by acting as a ‘‘mass storage’’ for a numerous catabolic genes. The presence of multiple catabolic pathways and gene homologs seems to be the basis of their catabolic versatility. However, many of the genes associated with the pathways are dispersed around the genome and it is becoming clear that their co-regulation is a feature of how the rhodoccci adapt to utilize many substrates.

1

Introduction

Members of the genus Rhodococcus are aerobic bacteria that possess the ability to degrade many recalcitrant and toxic pollutants in the environment and they have been discovered in many niches. Aspects of their phylogeny, physiology and biochemistry are reported elsewhere in the manual (cross reference: Volume 2(10) the Microbes) and will not be covered here. Instead this section will concentrate on the genetic basis of their catabolic abilities and developments regarding gene regulation and genomics; particularly related to catabolism and biodegradation. All of the rhodococci investigated to date appear to have large and complex genomes that may have acquired many genes by recombination in the distant past. There is increasing evidence that multiple pathways and gene homologs are present that further increases Rhodococcus catabolic versatility. Many also possess both large linear plasmids and circular plasmids that contribute greatly to the repertoire of catabolic genes. There appears to be an ability to adapt to degrade many new substrates, however the mechanisms underlying the flexibility of the Rhodococcus genome are not well understood. One striking feature is the presence of large linear plasmids in many of the strains isolated. Other characteristics are a system that promotes high frequency illegitimate recombination and the presence of relatively few transposons (although a small number of insertion sequences have been identified). Other, as yet unknown events of illegitimate recombination may serve to promote the introgression of DNA in their genomes without the help of mobile genetic elements (de Vries and Wackernagel, 2002). Aspects of their overall metabolic diversity and genetics have been covered previously in reviews (Bell et al., 1998; Gurtler et al., 2004; Kulakov and Larkin, 2002; Larkin et al., 2005; Larkin et al., 2006; McLeod and Eltis, 2008; O’Brien et al., 2002; van der Geize and Dijkhuizen, 2004; Warhurst and Fewson, 1994). This review will focus on some aspects of their genetic versatility and further emphasize the discoveries related to functional genomics.

2

The Genetic Basis of Biodegradation Capability

Despite an increased understanding of the wide range of catabolic abilities that the rhodococci possess, an understanding of their genetics and recombination mechanisms is still far from advanced. Despite Rhodococcus genetic diversity being immense, and hence the selection of a

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representative strain becomes difficult, recent advances in detailed analysis of genomic data is revealing many surprises. These observations are mostly based upon analysis of one strain; Rhodococcus jostii RHA1 whose genome has been extensively analyzed by Genome Canada and is publically available (see: http://www.rhodococcus.ca/). The genomes of other biodegradative strains; Rhodococcus aetherovorans strain I24 and Rhodococcus erythropolis strain PR4 have been analyzed but this data is not readily available and will not be the subject of discussion here. Equally, there is an ongoing genome sequence project for the animal pathogen Rhodococcus equi at the Sanger Centre, Cambridge, UK (see: http://www.sanger.ac.uk/Projects/ R_equi/) that will also not be addressed here. Experiments with RHA1 strain are providing the backbone of our current understanding in relation to functional genomics in the rhodococci. By way of a caveat, it is notable that a feature that can influence segregation of mutants and genetic elements, and which is often not considered, is their cellular pleomorphism. Many strains grow as short rods and cocci and also produce multinucleated filaments (Locci, 1984; Williams, 1976). RHA1 also displays this cellular morphology and this is an issue of note for subsequent studies and genetic analyses.

3

Genome Size and Composition

Bearing in mind the catabolic diversity of the rhodococi, it is not surprising that the genome sequence of Rhodococcus sp. strain RHA1, a polychlorinated-biphenyl (PCB)-degrading bacterium originally isolated on lindane as a substrate (Seto et al., 1995), is one of the largest bacterial genomes analyzed to date. It consists of 9,702,737 bp (with a G + C content of 67%) that is shared between three large linear plasmids; pRHL1 (1,100 kb), pRHL2 (450 kb) and pRHL3 (330 kb) and a linear chromosome (McLeod et al., 2006). In the case of each replicon, the telomeric sequences have been clearly identified. The genome sequence data is available for analysis at the Genome Canada site at: http://www.rhodococcus.ca/. The general conclusion is that RHA1 appears to have evolved to simultaneously catabolise a diverse range of organic compounds in an oxygen-rich environment. In this regard, it is particularly notable that there are at least 203 different oxygenases associated with the degradation pathways of aromatic compounds and steroids identified. In the case of aromatic compounds there appears to be at least 26 different ‘‘peripheral aromatic’’ pathways for a very wide range of compounds and 8 ‘‘central aromatic pathways.’’ In the light of this observation, it is surprising that the RHA1 genome appears to harbor few recent gene duplications and many genes also do not appear to have been acquired through recent horizontal transfer. However, in other Rhodococcus strains there is evidence that gene transfer is a key factor related to the degradation of many xenobiotic compounds as discussed elsewhere (See > Chapter 17, Vol. 3, Part 1). For example, there is clear evidence of transfer of genes associated with the catabolism of haloalkanes (Poelarends et al., 2000a, b), alkenes (Leahy et al., 2003), biphenyl (Taguchi et al., 2004, 2007), naphthalene and the explosive RDX (Seth-Smith et al., 2008) amongst various independently isolated strains. However, in the case of RHA1, it appears to have primarily acquired its large genome through more ancient gene duplications and gene transfers. It is notable that the RHA1 genome harbors only two intact insertion sequences and relatively few transposase genes (McLeod et al., 2006). The large genomes (over 7 Mb each) of two other rhodococci, Rhodococcus aetherovorans strain I24 and Rhodococcus erythropolis strain PR4, have also indicated the presence of multiple gene homologues and this is probably a feature of most of the bipodegradative rhodococci. However, the pathogen Rhodococcus equi has a

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smaller genome of just over 5 Mb. Further comparative analysis of these large genomes will no doubt reveal a wealth of catabolic genes and capacity to degrade many xenobiotic compounds.

4

Discovery of New Catabolic Pathways from the Genome Sequence

Because of the diverse catabolic genes present in RHA1, it is not surprising that analysis has led to the discovery of new pathways and abilities amongst the genome. Early analysis indicated a gene cluster that putatively encodes genes for taurine-pyruvate aminotransferase (Tpa) and alanine dehydrogenase (Ald) and for the catabolism and regulation of taurine catabolism. Experimentation confirmed growth on taurine and involvement of these genes (Denger et al., 2004). However, it is notable that these genes are separated from the associated, sulfoacetaldehyde acetyltransferase (xsc), phosphotransacetylase (pta) and possible ABC transporter (tauBC) genes. Proteomic analysis of RHA1 in response to growth on nitriles has led to the discovery of genes on the linear plasmid pRHL2; including a new class of acetonitrile hydratase (Okamoto and Eltis, 2007). Interestingly transcriptomic analysis of cells grown on cholesterol led to the discovery and confirmation of steroid catabolic genes that are also present in Mycobacterium tuberculosis (Van der Geize et al., 2007). Similar transcriptomic analyses have led to the conclusion that a propane monooxygenase (PrmA) is responsible for the degradation of the water contaminant N-nitrosodimethylamine in RHA1 (Sharp et al., 2007) and that phthalate is likely to be degraded solely via the protocatechuate pathway. However terephthalate is degraded via a bifurcated pathway that includes the catechol branch of the protocatechuate (Hara et al., 2007). These studies illustrate the hidden capabilities of RHA1 and the mechanism of co-regulation of these genes expressed from different genomic locations is complex and remains to be resolved and this issue will be explored further below.

5

The Role of Catabolic Plasmids

It is not surprising that cells of many of the rhodococci have been also shown to possess a wide variety of plasmid DNA molecules. These appear to range from small cryptic closed circular plasmids to the large linear plasmids as noted for the well characterized RHA1 strain noted above. Virtually every Rhodococcus strain analyzed to date appears to harbor plasmid DNA and linear plasmid DNA molecules have been detected in many strains. Although, the mechanisms of replication, or the ends of most of these molecules, have not been fully characterized, it was noted much earlier that in pHG207 (associated with hydrogen atrophy in Rhodococcus opacus) that the 50 imperfect inverted repeats were typical of actinomycete invertrons. (Kalkus et al., 1990, Kalkus et al., 1993, Kalkus et al., 1998). In RHA1, the sequence of pRHL3 has also been shown to be a typical actinomycete invertron, containing large terminal inverted repeats associated with a protein (Warren et al., 2004). Twenty one percent of its 300 putative genes, have a predicted catabolic function and are organized into three clusters. Four regions are likely to have been acquired by horizontal gene transfer involving transposition functions. There were similar observations for pBD2 from the isopropyl benzene utilizing

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strain Rhodococcus erythropolis BD2 (Stecker et al., 2003) where 23 of 99 putative genes probably have catabolic functions and 32 have possible transposition functions. There are many smaller circular plasmids noted in the rhodococci such as the cryptic plasmid pKA22 (4969 bp), pRTL1 (100 bp) encoding halolakane degradation genes (Kulakova et al., 1995) and a 150 bp a 150 kb plasmid in Rhodococcus sp. strain IGTS8 encoding genes that are involved in the desulphurisation of organosulphur compounds (Denis-Larose et al., 1997). However, the most interesting feature is the presence of many large linear plasmids. Such plasmids have been shown to encode genes for the catabolism of trichloroethene (Saeki et al., 1999), naphthalene (Kulakov et al., 2005; O’Brien et al., 2002; Uz et al., 2000), toluene (O’Brien et al., 2002; Priefert et al., 2004), alkylbenzene (Kim et al., 2002), biphenyl (Taguchi et al., 2004), chloroaromatic compounds (Konig et al., 2004). It is likely that multiple recombinations over a considerable time-span have resulted in many diverse genes becoming scattered around the genome that is also made up of several linear plasmids. It is notable that very few transposable elements have been characterised or observed (Kulakov et al., 1999; Lessard et al., 1999; McLeod et al., 2006; Nagy et al., 1997). Much earlier research however indicated that these bacteria possess mechanisms for a high frequency of illegitimate recombination (Kulakov and Larkin, 2002) and it can be postulated that this may mediate, alongside homologous recombination, the intro-regression of DNA without the need for many of mobile genetic elements as suggested for Acinetobacter (de Vries and Wackernagel, 2002). Indeed, earlier studies of plasmid integration in Rhodococcus fascians (Desomer et al., 1991) have indicated the involvement of a short palindromic sequence (CCGCGG) and that exogenous DNA bearing such a sequence could lead to a nonhomologous integration at a recombinational ‘‘hot-spot’’ sequence. Although this phenomenon has not been widely reported or studied it is possible that rhodococci may be capable of recombining easily with a wide variety of heterologous sequences.

6

Catabolic Gene Regulation and Expression

Studies of the regulation of a few Rhodococcus biodegradation gene clusters has revealed examples of both positive regulators (Komeda et al., 1996) and repressors (Barnes et al., 1997; Nga et al., 2004). More recently, studies based upon investigating the whole genome indicate that there is co-regulation of genetically unlinked transcriptional units and this is more intriguing. The most thoroughly investigated have been the multiple biphenyl/PCB degradation genes of RHA1 (also in Rhodococcus strain M5) that were originally noted as distributed in several clusters (Kitagawa et al., 2001; Yamada et al., 1998) on plasmids pRHL1 and pRHL2 (Shimizu et al., 2001). RHA1 expresses two biphenyl dioxygenases (BphA and EtbA/EbdA) when growing on biphenyl (Iwasaki et al., 2007) and the wider substrate range of the EtbA/EbdA dioxygenase suggested that it plays a more important role in the degradation of other compounds such as PCBs (Iwasaki et al., 2006). In Rhocococus M5, it has been demonstrated that a two-component regulatory system (bpdST ) regulates expression of some of the bph genes (Labbe et al., 1997) and this also appears to be the case for RHA1 (Takeda et al., 2004a, b) and expression of the o-xylene catabolic genes in Rhodococcus sp. strain DK17 (Kim et al., 2005). A putative GntR-like transcriptional regulation mechanism, involving narR1 and narR2, appears to be involved in several naphthalene degrading Rhodococcus strains (Kulakov et al., 2005). In this case there is also evidence that the catabolic genes are not organized into a single cluster and different

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strains have several homologous transcriptional units separated by non-homologous sequences containing direct and inverted repeats. This is further complicated by the demonstration of different promoter sequences initiating the expression of the homologous narAa – narB gene clusters and suggests that recombination events may be involved in the acquisition and alignment of regulatory regions with the catabolic genes. In Rhodococcus RHA1, as noted above, the degradation genes for benzoate (ben), phthalate (pad), uptake of phthalate (pat) and the genes for two branches of the b-ketoadipate pathway (cat and pca) are also dispersed. Indeed, some are contained on a putative ‘‘catabolic island’’ that is duplicated on plasmids pRHL1 and pRHL2. The regulatory interrelationship between the gene clusters is complex with involvement of pad and pat gene products in phthalate degradation and ben and cat gene products in benzoate degradation. Expression of the pca products are also implicated as they are present after growth on both substrates (Patrauchan et al., 2005). To further elucidate the complex expression of such genes, both proteomic and gene disruption techniques have been used to investigate the degradation of benzene, styrene, biphenyl, and ethylbenzene in RHA1. Cells grown on biphenyl, ethylbenzene, or benzene produced enzymes associated with both biphenyl and ethyl benzene catabolism and enzymes from at least two sets of lower biphenyl pathways. Styrene-grown cells contained no ethylbezene pathway enzymes and only one set of lower biphenyl pathway enzymes. Biphenyl dioxygenase was essential for growth on benzene or styrene but the ethylbenzene dioxygenase was not required for growth on any of the substrates (Patrauchan et al., 2008). This indicates that co-expression of genes from different loci may be commonplace. Indeed, the utilization of phenylacetic acid is encoded in part by 13 paa genes on chromosome. A single transcript encodes 11 genes but production of a further 146 proteins was induced by growth on phenylacetic acid (Navarro-Llorens et al., 2005). Further evidence for such co-expression has arisen from analysis of the RHA1 transcriptome using various substrates. RHA1 encodes multiple isoenzymes for most of the steps in biphenyl catabolism. Co-expression of these is clearly associated with the catabolism of not only biphenyl, but also ethylbenzene and PCBs. Transcriptomic analysis of over 8000 potential catabolic genes indicate that for biphenyl and ethylbenzene, over 320 are up-regulated, unlike benzoate that led to up-regulation of 65 genes. There was no difference in the expression of key catabolic genes for ethylbenzene and biphenyl indicating the likelihood of a common regulation for these (Goncalves et al., 2006).

7

Gene Expression in Response to Environmental Stresses

To date there have been few studies that have addressed how rhodococci respond to stresses associated with the general environment. It has been acknowledged that these bacteria can withstand desiccation and that this could be a key factor in their survival in such environments. Interestingly, a transcriptomic study indicated that those genes associated with desiccation were maximally up-regulated upon complete drying of the cells. These included putative genes associated with oxidative stress, biosynthesis of the compatible solute ectoine and sigma factors indicating their putative role in regulation (LeBlanc et al., 2008). In terms of expression in the environment there has been little research, however it has been shown that genes associated with biphenyl degradation are expressed in biphenyl amended soils (Wang et al., 2008).

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Research Needs

Analysis of the biochemical diversity, physiology and the genomes of Rhodococcus strains involved in the biodegradation of pollutants is slowly revealing why they can adapt to catabolise many different substrates. However, whole genome analysis is based largely upon one strain, Rhodococcus jostii RHA1 that indicates that they have acquired many copies of genes in very large genomes that include large linear plasmids. There is a need now to develop whole genome analysis of other strains and investigate further how recombination and gene transfer occurs. The research to date goes some way to explain how single strains can utilize a remarkably wide range of substrates. The surprise is the extent to which the pathways for biodegradation are dispersed around the genome and apparently the subject of co-regulation. The elucidation of how this occurs is the main challenge in understanding rhodococci and their relationship with the polluted environment. Also of importance is demonstrating the range of genes expressed by rhodococci in the environment of polluted soils and waters where remediation occurs. This would go some way to defining the optimal niches for these bacteria and maximize their potential for bioremediation.

References Barnes MR, Duetz WA, Williams PA (1997) A 3-(3hydroxyphenyl)propionic acid catabolic pathway in Rhodococcus globerulus PWD1: cloning and characterization of the hpp operon. J Bacteriol 179: 6145–6153. Bell KS, Philp JC, Aw DW, Christofi N (1998) The genus Rhodococcus. J Appl Microbiol 85: 195–210. de Vries J, Wackernagel W (2002) Integration of foreign DNA during natural transformation of Acinetobacter sp. by homology-facilitated illegitimate recombination. Proc Natl Acad Sci USA 99: 2094–2099. Denger K, Ruff J, Schleheck D, Cook AM (2004) Rhodococcus opacus expresses the xsc gene to utilize taurine as a carbon source or as a nitrogen source but not as a sulfur source. Microbiology 150: 1859–1867. Denis-Larose C, Labbe D, Bergeron H, Jones AM, Greer CW, al-Hawari J, Grossman MJ, Sankey BM, Lau PC (1997) Conservation of plasmid-encoded dibenzothiophene desulfurization genes in several rhodococci. Appl Environ Microbiol 63: 2915–2919. Desomer J, Crespi M, Van Montagu M (1991) Illegitimate integration of non-replicative vectors in the genome of Rhodococcus fascians upon electrotransformation as an insertional mutagenesis system. Mol Microbiol 5: 2115–2124. Goncalves ER, Hara H, Miyazawa D, Davies JE, Eltis LD, Mohn WW (2006) Transcriptomic assessment of isozymes in the biphenyl pathway of Rhodococcus sp. strain RHA1. Appl Environ Microbiol 72: 6183–6193.

Gurtler V, Mayall BC, Seviour R (2004) Can whole genome analysis refine the taxonomy of the genus Rhodococcus? FEMS Microbiol Rev 28: 377–403. Hara H, Eltis LD, Davies JE, Mohn WW (2007) Transcriptomic analysis reveals a bifurcated terephthalate degradation pathway in Rhodococcus sp. strain RHA1. J Bacteriol 189: 1641–1647. Iwasaki T, Miyauchi K, Masai E, Fukuda M (2006) Multiple-subunit genes of the aromatic-ringhydroxylating dioxygenase play an active role in biphenyl and polychlorinated biphenyl degradation in Rhodococcus sp. strain RHA1. Appl Environ Microbiol 72: 5396–5402. Iwasaki T, Takeda H, Miyauchi K, Yamada T, Masai E, Fukuda M (2007) Characterization of two biphenyl dioxygenases for biphenyl/PCB degradation in A PCB degrader, Rhodococcus sp. strain RHA1. Biosci Biotechnol Biochem 71: 993–1002. Kalkus J, Dorrie C, Fischer D, Reh M, Schlegel HG (1993) The giant linear plasmid pHG207 from Rhodococcus sp. encoding hydrogen autotrophy: characterization of the plasmid and its termini. J Gen Microbiol 139: 2055–2065. Kalkus J, Menne R, Reh M, Schlegel HG (1998) The terminal structures of linear plasmids from Rhodococcus opacus. Microbiology 144(Pt 5): 1271–1279. Kalkus J, Reh M, Schlegel HG (1990) Hydrogen autotrophy of Nocardia opaca strains is encoded by linear megaplasmids. J Gen Microbiol 136: 1145–1151. Kim D, Chae JC, Zylstra GJ, Sohn HY, Kwon GS, Kim E (2005) Identification of two-component regulatory

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genes involved in o-xylene degradation by Rhodococcus sp. strain DK17. J Microbiol 43: 49–53. Kim D, Kim YS, Kim SK, Kim SW, Zylstra GJ, Kim YM, Kim E (2002) Monocyclic aromatic hydrocarbon degradation by Rhodococcus sp. strain DK17. Appl Environ Microbiol 68: 3270–3278. Kitagawa W, Miyauchi K, Masai E, Fukuda M (2001) Cloning and characterization of benzoate catabolic genes in the gram-positive polychlorinated biphenyl degrader Rhodococcus sp. strain RHA1. J Bacteriol 183: 6598–6606. Komeda H, Hori Y, Kobayashi M, Shimizu S (1996) Transcriptional regulation of the Rhodococcus rhodochrous J1 nitA gene encoding a nitrilase. Proc Natl Acad Sci USA 93: 10572–10577. Konig C, Eulberg D, Groning J, Lakner S, Seibert V, Kaschabek SR, Schlomann M (2004) A linear megaplasmid, p1CP, carrying the genes for chlorocatechol catabolism of Rhodococcus opacus 1CP. Microbiology 150: 3075–3087. Kulakov LA, Chen S, Allen CC, Larkin MJ (2005) Webtype evolution of rhodococcus gene clusters associated with utilization of naphthalene. Appl Environ Microbiol 71: 1754–1764. Kulakov LA, Larkin MJ (2002) Genomic organization of Rhodococcus. In A Danchin (ed.). Genomics of GC-Rich gram-positive bacteria. Norfolk UK: Caister Academic Press, pp. 15–46. Kulakov LA, Poelarends GJ, Janssen DB, Larkin MJ (1999) Characterization of IS2112, a new insertion sequence from Rhodococcus, and its relationship with mobile elements belonging to the IS110 family. Microbiology 145(Pt 3): 561–568. Kulakova AN, Stafford TM, Larkin MJ, Kulakov LA (1995) Plasmid pRTL1 controlling 1-chloroalkane degradation by Rhodococcus rhodochrous NCIMB 13064. Plasmid 33: 208–217. Labbe D, Garnon J, Lau PC (1997) Characterization of the genes encoding a receptor-like histidine kinase and a cognate response regulator from a biphenyl/ polychlorobiphenyl-degrading bacterium, Rhodococcus sp. strain M5. J Bacteriol 179: 2772–2776. Larkin MJ, Allen CCR, Kulakov LA (2006) Biodegradation by Members of the Genus Rhodococcus: Biochemistry, Physiology,and Genetic Adaptation. Adv Appl Microbiol 59: 1–28. Larkin MJ, Kulakov LA, Allen CCR (2005) Biodegradation and Rhodococcus - masters of catabolic versatility. Curr Opin Biotechnol 16(3): 282–290. Leahy JG, Batchelor PJ, Morcomb SM (2003) Evolution of the soluble diiron monooxygenases. FEMS Microbiol Rev 27: 449–479. LeBlanc JC, Goncalves ER, Mohn WW (2008) Global response to desiccation stress in the soil actinomycete Rhodococcus jostii RHA1. Appl Environ Microbiol 74: 2627–2636.

Lessard PA, O’Brien XM, Ahlgren NA, Ribich SA, Sinskey AJ (1999) Characterization of IS1676 from Rhodococcus erythropolis SQ1. Appl Microbiol Biotechnol 52: 811–819. Locci R (1984) Morphology. In: The Biology of the Actinomycetes. M Goodfellow (ed.). New York: Academic press, pp. 165–199. McLeod MP, Eltis LD (2008) Genomic insights into the aerobic pathways for degradation of organic pollutants. In E Diaz (ed.). Microbial Biodegradation: Genomics and Molecular Biology, 1st edn. UK: Caister Academic Press. McLeod MP, Warren RL, Hsiao WW, Araki N, Myhre M, Fernandes C, Miyazawa D, Wong W, Lillquist AL, Wang D, Dosanjh M, Hara H, Petrescu A, Morin RD, Yang G, Stott JM, Schein JE, Shin H, Smailus D, Siddiqui AS, Marra MA, Jones SJ, Holt R, Brinkman FS, Miyauchi K, Fukuda M, Davies JE, Mohn WW, Eltis LD (2006) The complete genome of Rhodococcus sp. RHA1 provides insights into a catabolic powerhouse. Proc Natl Acad Sci USA 103: 15582–15587. Nagy I, Schoofs G, Vanderleyden J, De Mot R (1997) Transposition of the IS21-related element IS1415 in Rhodococcus erythropolis. J Bacteriol 179: 4635–4638. Navarro-Llorens JM, Patrauchan MA, Stewart GR, Davies JE, Eltis LD, Mohn WW (2005) Phenylacetate catabolism in Rhodococcus sp strain RHA1: a central pathway for degradation of aromatic compounds. J Bacteriol 187: 4497–4504. Nga DP, Altenbuchner J, Heiss GS. (2004) NpdR, a repressor involved in 2,4,6-trinitrophenol degradation in Rhodococcus opacus HL PM-1. J Bacteriol 186: 98–103. O’Brien XM, Parker JA, Lessard PA, Sinskey AJ (2002) Engineering an indene bioconversion process for the production of cis-aminoindanol: a model system for the production of chiral synthons. Appl Microbiol Biotechnol 59: 389–399. Okamoto S, Eltis LD (2007) Purification and characterization of a novel nitrile hydratase from Rhodococcus sp. RHA1. Mol Microbiol 65: 828–838. Patrauchan MA, Florizone C, Dosanjh M, Mohn WW, Davies J, Eltis LD (2005) Catabolism of benzoate and phthalate in Rhodococcus sp. strain RHA1: redundancies and convergence. J Bacteriol 187: 4050–4063. Patrauchan MA, Florizone C, Eapen S, Gomez-Gil L, Sethuraman B, Fukuda M, Davies J, Mohn WW, Eltis LD (2008) Roles of ring-hydroxylating dioxygenases in styrene and benzene catabolism in Rhodococcus jostii RHA1. J Bacteriol 190: 37–47. Poelarends GJ, Kulakov LA, Larkin MJ, van Hylckama Vlieg JE, Janssen DB (2000a) Roles of horizontal gene transfer and gene integration in evolution

Rhodococcus: Genetics and Functional Genomics of 1,3-dichloropropene- and 1,2-dibromoethanedegradative pathways. J Bacteriol 182: 2191–2199. Poelarends GJ, Zandstra M, Bosma T, Kulakov LA, Larkin MJ, Marchesi JR, Weightman AJ, Janssen DB (2000b) Haloalkane-utilizing Rhodococcus strains isolated from geographically distinct locations possess a highly conserved gene cluster encoding haloalkane catabolism. J Bacteriol 182: 2725–2731. Priefert H, O’Brien XM, Lessard PA, Dexter AF, Choi EE, Tomic S, Nagpal G, Cho JJ, Agosto M, Yang L, Treadway SL, Tamashiro L, Wallace M, Sinskey AJ (2004) Indene bioconversion by a toluene inducible dioxygenase of Rhodococcus sp. I24. Appl Microbiol Biotechnol 65: 168–176. Saeki H, Akira M, Furuhashi K, Averhoff B, Gottschalk G (1999) Degradation of trichloroethene by a linear-plasmid-encoded alkene monooxygenase in Rhodococcus corallinus (Nocardia corallina) B-276. Microbiology 145(Pt 7): 1721–1730. Seth-Smith HM, Edwards J, Rosser SJ, Rathbone DA, Bruce NC (2008) The explosive-degrading cytochrome P450 system is highly conserved among strains of Rhodococcus spp. Appl Environ Microbiol 74: 4550–4552. Seto M, Masai E, Ida M, Hatta T, Kimbara K, Fukuda M, Yano K (1995) Multiple polychlorinated biphenyl transformation systems in the Gram-Positive Bacterium Rhodococcus sp. Strain RHA1. Appl Environ Microbiol 61: 4510–4513. Sharp JO, Sales CM, LeBlanc JC, Liu J, Wood TK, Eltis LD, Mohn WW, Alvarez-Cohen L (2007) An inducible propane monooxygenase is responsible for N-nitrosodimethylamine degradation by Rhodococcus sp. strain RHA1. Appl Environ Microbiol 73: 6930–6938. Shimizu S, Kobayashi H, Masai E, Fukuda M (2001) Characterization of the 450-kb linear plasmid in a polychlorinated biphenyl degrader, Rhodococcus sp. strain RHA1. Appl Environ Microbiol 67: 2021–2028. Stecker C, Johann A, Herzberg C, Averhoff B, Gottschalk G (2003) Complete nucleotide sequence and genetic organization of the 210-kilobase linear plasmid of Rhodococcus erythropolis BD2. J Bacteriol 185: 5269–5274. Taguchi K, Motoyama M, Iida T, Kudo T (2007) Polychlorinated biphenyl/biphenyl degrading gene clusters in Rhodococcus sp. K37, HA99, and TA431 are different from well-known bph gene clusters of Rhodococci. Biosci Biotechnol Biochem 71: 1136–1144. Taguchi K, Motoyama M, Kudo T (2004) Multiplicity of 2,3-dihydroxybiphenyl dioxygenase genes in the Gram-positive polychlorinated biphenyl degrading

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bacterium Rhodococcus rhodochrous K37. Biosci Biotechnol Biochem 68: 787–795. Takeda H, Hara N, Sakai M, Yamada A, Miyauchi K, Masai E, Fukuda M (2004a) Biphenyl-inducible promoters in a polychlorinated biphenyl-degrading bacterium, Rhodococcus sp. RHA1. Biosci Biotechnol Biochem 68: 1249–1258. Takeda H, Yamada A, Miyauchi K, Masai E, Fukuda M (2004b) Characterization of transcriptional regulatory genes for biphenyl degradation in Rhodococcus sp. strain RHA1. J Bacteriol 186: 2134–2146. Uz I, Duan YP, Ogram A (2000) Characterization of the naphthalene-degrading bacterium, Rhodococcus opacus M213. FEMS Microbiol Lett 185: 231–238. van der Geize R, Dijkhuizen L (2004) Harnessing the catabolic diversity of rhodococci for environmental and biotechnological applications. Curr Opin Microbiol 7: 255–261. Van der Geize R, Yam K, Heuser T, Wilbrink MH, Hara H, Anderton MC, Sim E, Dijkhuizen L, Davies JE, Mohn WW, Eltis LD (2007) A gene cluster encoding cholesterol catabolism in a soil actinomycete provides insight into Mycobacterium tuberculosis survival in macrophages. Proc Natl Acad Sci USA 104: 1947–1952. Wang Y, Shimodaira J, Miyasaka T, Morimoto S, Oomori T, Ogawa N, Fukuda M, Fujii T (2008) Detection of bphAa gene expression of Rhodococcus sp. strain RHA1 in soil using a new method of RNA preparation from soil. Biosci Biotechnol Biochem 72: 694–701. Warhurst AM, Fewson CA (1994) Biotransformations catalyzed by the genus Rhodococcus. Crit Rev Biotechnol 14: 29–73. Warren R, Hsiao WW, Kudo H, Myhre M, Dosanjh M, Petrescu A, Kobayashi H, Shimizu S, Miyauchi K, Masai E, Yang G, Stott JM, Schein JE, Shin H, Khattra J, Smailus D, Butterfield YS, Siddiqui A, Holt R, Marra MA, Jones SJ, Mohn WW, Brinkman FS, Fukuda M, Davies J, Eltis LD (2004) Functional characterization of a catabolic plasmid from polychlorinated- biphenyl-degrading Rhodococcus sp. strain RHA1. J Bacteriol 186: 7783–7795. Williams ST (1976) The micromorpholpgy and fine structure of nocardioformorganisms. In: The Biology of the Nocardiae. M Goodfellow, GH Brownell, JA Serrano (eds.). New York: Academic Press pp. 103–140. Yamada A, Kishi H, Sugiyama K, Hatta T, Nakamura K, Masai E, Fukuda M (1998) Two nearly identical aromatic compound hydrolase genes in a strong polychlorinated biphenyl degrader, Rhodococcus sp. strain RHA1. Appl Environ Microbiol 64: 2006–2012.

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39 Phylogenomics of Aerobic Bacterial Degradation of Aromatics D. Pe´rez-Pantoja1 . R. Donoso1 . H. Junca2,3 . B. Gonza´lez1 . D. H. Pieper3 Departamento de Gene´tica Molecular y Microbiologı´a, Facultad de Ciencias Biolo´gicas, NM-EMBA, CASEB, P. Universidad Cato´lica de Chile, Santiago, Chile 2 Centro Colombiano de Metageno´mica y Bioinforma´tica de Ambientes Extremos (GeBiX) – Grupo de Gene´tica Molecular, Corporacio´n CorpoGen, Bogota´, Colombia 3 Biodegradation Research Group, Department of Microbial Pathogenesis, HZI – Helmholtz Centre for Infection Research, Braunschweig, Germany [email protected] 1

1

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1356

2

Aerobic Aromatic Catabolic Routes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1356

3

Sequenced Bacterial Genomes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1357

4 4.1 4.2 4.3 4.4 4.5 4.6 4.7 4.8

Spread of Members of Gene Families . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1359 Intradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1359 EXDO I Family . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1361 Lig B Superfamily . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1365 Cupin Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1367 Other Extradiol Dioxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1368 Diiron Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1368 Flavoprotein Monooxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1369 Rieske Non-Heme Iron Oxygenases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1375

5 5.1 5.2 5.3 5.4 5.5 5.6

Metabolism Diversity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1378 Metabolism by Bacteria Outside the Actinobacterial and Proteobacterial Phyla . . . . . . 1378 Actinobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1382 Proteobacteria . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1383 Pathway Redundancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1388 Gene Redundancy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1389 Superbugs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1390

6

Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1391

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_95, # Springer-Verlag Berlin Heidelberg, 2010

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Phylogenomics of Aerobic Bacterial Degradation of Aromatics

Abstract: Aromatic compounds are widely distributed in nature. They are found as lignin components, aromatic amino acids, and xenobiotic compounds, among others. Microorganisms, mostly bacteria, degrade an impressive variety of such chemical structures. Various aerobic aromatic catabolic pathways have been reported in bacteria, which typically consist of activation of the aromatic ring through oxygenases or CoA ligases and ring cleavage of di- or trihydroxylated intermediates or dearomatized CoA derivatives. We survey almost 900 sequenced bacterial genomes for the presence of genes encoding key enzymes of aromatic metabolic pathways, including ring-cleavage enzymes as well as enzymes activating aromatics or dearomatizing CoA derivatives. The metabolic diversity is discussed from two angles: the spread of such key activities among different bacterial phyla and the overall metabolic potential of members of bacterial genera.

1

Introduction

A few non-mutually exclusive choices are possible to address the analysis of the genetic basis of bacterial degradation of aromatic compounds. One is to select a few well-studied bacterial catabolic models and go in depth into their genetic organization of aromatic catabolism genes (Jimenez et al., 2002; Pe´rez-Pantoja et al., 2008). Another approach is to select a few central catabolic pathways and to assess the similarities and differences in gene organization, substrate range, and regulatory elements, among the bacteria where such pathways have been described. A third possibility is to look for all the aromatic catabolism pathways present in bacteria, searching in the growing database of sequenced bacterial genomes. The latter, by definition, is a less in-depth analysis but has the broader coverage possible today. We selected the latter approach, because we think it provides clues on the distribution of catabolic properties among bacterial phyla, gives some hints on the ecological functions of specific bacterial groups, defines underscored research objectives, and gives a better overview of the genetic basis of bacterial catabolism of aromatics. The phylogenomic approach to study the organization of aromatic degradation is based on the selection of sequences of key catabolic functions to fish into the sequenced genome database, followed by refinement of the positive scores. With this information, the genomes can be analyzed in terms of presence/absence of catabolic abilities among bacterial groups, new enzyme families based on the sequence similarity be defined, new putative functions be suggested, and evolutionary links among different groups of sequences be addressed. Of course such approach has some limitations, as most of the new data are not supported by biochemical or genetic studies. To minimize such limitations, the selected sequence probes were derived from both biochemical and genetic well-studied systems. One of the main purposes of the following material is to provide to the reader new research venues to get a deeper knowledge on bacterial catabolism of aromatics.

2

Aerobic Aromatic Catabolic Routes

Bacterial degradation of aromatic compounds and their haloaromatic derivatives has been well studied (See > Chapter 4, Vol. 2, Part 2; > Chapter 5, Vol. 2, Part 2). Various pathways for degradation of these compounds by bacteria have been reported. The activation of the

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

aromatic ring commonly proceeds by members of one of three superfamilies: the Rieske nonheme iron oxygenases usually catalyzing the incorporation of two oxygen atoms (although some members of this superfamily also catalyze monooxygenations) (Gibson and Parales, 2000), the flavoprotein monooxygenases (van Berkel et al., 2006), and the soluble diiron multicomponent oxygenases (Leahy et al., 2003). Further metabolism is achieved through di- or trihydroxylated aromatic intermediates. Alternatively, activation is mediated by CoA ligases and the formed CoA derivatives are subjected to oxygenations. This can proceed through 2-aminobenzoyl-CoA monooxygenase/reductase, an enzyme that catalyzes both monooxygenation and hydrogenation, and where the N-terminal part of the protein shows similarities to single-component flavin monooxygenases (Buder and Fuchs, 1989). Alternatively, the aromatic CoA derivative is attacked by multicomponent enzymes, where the oxygenase subunits belong to the diiron oxygenases, like in phenylacetyl-CoA (Ismail et al., 2003) or benzoyl-CoA oxygenase (Zaar et al., 2004). Various further key reactions channeling aromatics to central di- or trihydroxylated intermediates, such as the processing of side chains or demethylations, will not be discussed here (See > Chapter 4, Vol. 2, Part 2). The further aerobic degradation of di- or trihydroxylated intermediates can be catalyzed by either intradiol or extradiol dioxygenases. While all intradiol dioxygenases described thus far belong to the same superfamily, members of at least three different families are reported to be involved in the extradiol ring cleavage of hydroxylated aromatics. Type I extradiol dioxygenases (e.g., catechol 2,3-dioxygenases) belong to the vicinal oxygen chelate superfamily enzymes (Gerlt and Babbitt, 2001), the type II or LigB superfamily of extradiol dioxygenases which comprise among other protocatechuate 4,5-dioxygenases (Sugimoto et al., 1999) and the type III enzymes such as gentisate dioxygenases which comprise enzymes belonging to the cupin superfamily (Dunwell et al., 2000). However, even though belonging to different families, all three types of extradiol dioxygenases share similar active sites and all type I, type II, and various type III enzymes have the same iron ligands, two histidine and one glutamate, that constitute the 2-His 1-carboxylate structural motif. The recently identified benzoquinol 1,2-dioxygenase from the 4-hydroxyacetophenone-degrading Pseudomonas fluorescens ACB that displays no significant sequence identity with known dioxygenases may constitute the prototype of a novel fourth class of Fe2+-dependent dioxygenases (Moonen et al., 2008).

3

Sequenced Bacterial Genomes

Currently (as of September 2008) approximately 1,000 genomes have been sequenced and three quarters of them finished. For the purpose of this review, we concentrated on genomes that were simultaneously represented in both the Integrated Microbial Genomes (IMG) database at DOE Joint Genome Institute (JGI) (img.jgi.doe.gov/cgi-bin/pub/main.cgi?page= home) and the National Center for Biotechnology Information (NCBI) database at National Institute of Health (NIH) (www.ncbi.nlm.nih.gov/sutils/genom_table.cgi), summing up to 822 genomes. The number of representatives of the bacterial phyla in these public databases is highly variable: from a very few members from the phyla Aquificae (2) Acidobacteria (2), Chlamydiae (11), Chlorobi (10), Chloroflexi (8), Deinococcus/Thermus (4), Fusobacteria (2), Lentisphaerae (2), Planctomycetes (3), Spirochaetes (9), Thermotogae (6), and Verrucomicrobia (1); the medium represented phyla: Actinobacteria (53), Bacteroidetes (28), Cyanobacteria (40), and the Proteobacteriales d- (23) and e- classes (28); and the highly represented phylum Firmicutes (182) and the a- (112) b- (71) and g- (223) classes of Proteobacteria (besides two

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Phylogenomics of Aerobic Bacterial Degradation of Aromatics

. Figure 1 (Continued)

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Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

unclassified Proteobacteria). Despite of that, the number of bacterial genomes is now significant to search for the presence/absence of the main catabolic pathways for aromatic compounds to provide a reasonable idea about the spread of these catabolic abilities among the main phylogenetic groups.

4

Spread of Members of Gene Families

4.1

Intradiol Dioxygenases

The intradiol cleavage of catechol to muconate and of protocatechuate to 3-carboxymuconate by catechol 1,2-dioxygenases and protocatechuate 3,4-dioxygenases, respectively, is a central reaction in the metabolism of various aromatic compounds (> Fig. 1). Hydroxybenzoquinol (1,2,4-trihydroxybenzene) is also a central intermediate in the degradation of a variety of aromatic compounds such as resorcinol (> Fig. 1), with hydroxybenzoquinol 1,2-dioxygenase as key enzyme, catalyzing intradiol cleavage to form 3-hydroxy-cis,cis-muconate and its tautomer, maleylacetate. Among the different groups of enzymes significant metabolic crossreactivity is usually not observed. Phylogenetic analysis of the deduced protein sequences of intradiol dioxygenases encoded in the genomes of bacteria sequenced so far showed the presence of seven clusters as indicated in > Fig. 2. Based on biochemical or genetically validated representatives, cluster 1 comprises hydroxybenzoquinol dioxygenases, cluster 2 proteobacterial catechol 1,2-dioxygenases, cluster 3 actinobacterial catechol 1,2-dioxygenases, and clusters 5 and 7 the a- and b-subunits of protocatechuate 3,4-dioxygenases, respectively. Enzymes of cluster 6 are obviously related to the b-subunits of protocatechuate dioxygenases, however, in no case genes encoding these enzymes are clustered with genes encoding putative a- subunits, and the function of these enzymes remains to be elucidated. Similarly, the function of enzymes of cluster 4 wait for clarification. Intradiol dioxygenases are nearly exclusively found in two phyla, the Actinobacteria and the Proteobacteria. However, protocatechuate 3,4-dioxygenases were observed in one of the two sequenced Deinococci, i.e., Deinococcus geothermalis DSM 11300 and one of the two sequenced Acidobacteria, i.e., Solibacter usitatus Ellin6076. Considering the wide spread of Acidobacteria in the environment, their involvement in aromatic degradation under natural

. Figure 1 Aerobic metabolism of aromatics via di- or trihydroxylated intermediates, or via CoA derivatives. Peripheral hydroxylation reactions can be catalyzed by flavoprotein monooxygenases, Rieske non-heme iron oxygenases or soluble diiron oxygenases. Alternatively, aromatics can be activated through CoA ligases followed by dearomatization catalyzed by members of the flavoprotein monooxygenases or soluble diiron oxygenases. 4-Hydroxyphenylpyruvate dioxygenase is indicated by a Central di- or trihydroxylated intermediates are subjected to ring cleavage by intradiol dioxygenases or extradiol dioxygenases of the vicinal chelate superfamily, the LigB superfamily or the cupin superfamily. Ring-cleavage products are channeled to the Krebs cycle via central reactions.

1359

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Phylogenomics of Aerobic Bacterial Degradation of Aromatics

. Figure 2 Evolutionary relationships among intradiol dioxygenases. The evolutionary history was inferred using the neighbor joining method after alignment of sequences using MUSCLE (Edgar, 2004). All positions containing alignment gaps and missing data were eliminated only in pairwise sequence comparisons. Wedges represent enzyme clusters as described in the text. Deduced protein sequences not falling inside the defined clusters are also indicated. Wedge length is a measure of evolutionary distance from the common ancestor. Phylogenetic analyses were conducted in MEGA4 (Tamura et al., 2007). Cluster 1 comprises hydroxybenzoquinol dioxygenases, cluster 2 proteobacterial catechol 1,2-dioxygenases, cluster 3 actinobacterial catechol 1,2-dioxygenases, and clusters 5 and 7 the a- and b-subunits of protocatechuate 3,4-dioxygenases, respectively. The functions of enzymes of clusters 4 and 6 remain to be elucidated.

conditions has to be considered. Actually, Acidobacteria have been implied to be involved in the biogeochemical cycles of rhizosphere soil (Lee et al., 2008). Regarding catechol 1,2-dioxygenases, where two lineages have previously been described (Eulberg et al., 1997), phylogenetic analysis confirmed that cluster 3 enzymes are restricted to members of the order Actinomycetales of the Actinobacteria, and catechol intradiol cleavage pathways were observed in the majority of Corynebacteria, Arthrobacter, Mycobacteria, and Nocardiaceae. Usually, Actinobacteria possessing a catechol intradiol cleavage pathway also harbor a protocatechuate intradiol cleavage. However, Streptomyces strains seem to be

39

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

endowed only with the protocatechuate branch. A hydroxybenzoquinol pathway seems to be spread only in Corynebacteria and out of the Mycobacteria, only Mycobacterium smegmatis and M. vanbaalenii are endowed with such a pathway. As shown in > Table 1, intradiol dioxygenases can be identified in 11 out of 19 a-proteobacterial, 2 out of 10 b-proteobacterial, and 4 out of 29 g-proteobacterial families and are absent in d- or e- proteobacteria. Significant differences in gene spread were observed among families. Catechol intradiol pathways are observed in nearly all Pseudomonas strains and are absent only from the genomes of P. syringae and P. mendocina. The last one is also the only Pseudomonas strain devoid of a protocatechuate intradiol pathway. Similarly, both protocatechuate and catechol pathways are observed in all Burkholderia genomes. Interestingly, catechol intradiol cleavage pathways were only exceptionally observed in a-Proteobacteria. In contrast, a catechol pathway is absent in Rhizobiaceae, which, however, often bear a hydroxybenzoquinol pathway. Also Bradyrhizobiaceae, none of which has a catechol pathway, are usually endowed with a hydroxybenzoquinol pathway except for Nitrobacter strains.

4.2

EXDO I Family

The extradiol ring cleavage of catechol is typically catalyzed by type I extradiol dioxygenases (EXDO I), which belong to the vicinal oxygen chelate superfamily (Gerlt and Babbitt, 2001). . Table 1 Intradiol dioxygenases observed in genomes of Proteobacteria

Class

Family

Protocatechuate 3,4-dioxygenase (Pca34)

a

Caulobacteraceae (2)

a

Aurantimonadaceae (2)

+

a

Bradyrhizobiaceae (11)

( )

a

Brucellaceae (6)

++

Catechol 1,2-dioxygenase (Cat12)

Hydroxybenzo-quinol dioxygenase (Hqu)

++

a

Methylobacteriaceae (3)

+

a

Phyllobactriaceae (3)

++

a

Rhizobiaceae (6)

++

a

Rhodobacteraceae (24)

++

a

Xanthobacteriaceae (2)

++

a

Acetobacteraceae (3)

a

Sphingomonadaceae (5)

b

++ + ++ + +

++

( )

+

( )

Burkholderiaceae (43)

++

++

+

b

Comamonadaceae (8)

( )

+

+

g

Oceanosprillaceae (3)

+

+

g

Moraxellaceae (5)

+

+

g

Pseudomonadaceae (19)

++

++

g

Xanthomonadaceae (11)

+

+

( )

++; More than 60% of the sequenced genomes of these bacterial taxa comprise a gene encoding the mentioned activity (number of sequenced representatives is given in parentheses); +, between 20 and 60%; ( ), less than 20%; , not observed

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The EXDO I family comprises enzymes that catalyze the dioxygenolytic ring fission of the catecholic derivatives in several bacterial mono- and polyaromatics biodegradation pathways (Eltis and Bolin, 1996) (> Fig. 1) like those involved in degradation of benzene, toluene, phenol, biphenyl, naphthalene, dibenzofuran, 4-hydroxyphenylacetate, p-cymene, or diterpenoid compounds such as abietate. They catalyze the meta-cleavage of catechol to 2-hydroxymuconic semialdehyde (catechol 2,3-dioxygenases, C23O), of 2,3-dihydroxybiphenyl (2,3-dihydroxybiphenyl 1,2-dioxygenases, BphC), 1,2-dihydroxynaphthalene (NahC), homoprotocatechuate (homoprotocatechuate 2,3-dioxygenases, HpaD), 2,3-dihydroxyp-cumate (2,3-dihydroxy-p-cumate-3,4-dioxygenases CmtC), and 7-oxo-11,12-dihydroxydehydroabietate (DitC), among others (see also > Fig. 1). In many cases, the respective genes are localized in catabolic pathway gene clusters such that their actual function can easily be deduced. However, in various cases multiple EXDO I activities are observed in a single strain and often their function remains unproven (Maeda et al., 1995). Here, the names are given to the enzymes according to the preferential activity observed, but in many cases there is a range of structurally similar substrates that can be metabolized by the same enzyme with varying catalytic efficacies and the ‘‘natural’’ substrate has not yet been identified. Because genome annotations pipelines are in many cases using the NCBI Conserved Domains Database (CDD), which is in turn, interconnected with the Wellcome Trust Sanger Institute Pfam database descriptions, all EXDO I genes found in the genome sequences are recognized and annotated with the superfamily name as Glyoxalase/bleomycin resistance protein/dioxygenase (InterPro: IPR004360, pfam00903: Glyoxalase). However, in the majority of cases, a more precise annotation of several genomic sequences as EXDO I would be possible, as they show conservation of the Prosite PS00082 extradiol ring-cleavage dioxygenases signature [GNTIV]-x-H-x(5,7)-[LIVMF]-Y-x(2)-[DENTA]-P-x-[GP]-x(2,3)-E. Phylogenetic analysis of the deduced protein sequences of EXDO I encoded in the genomes of bacteria sequenced so far, and retrieved after iterative PSI Blast searches using representative proteins of major clusters where a function has been described as seeds show the presence of three major evolutionary lineages (> Fig. 3). One of these lineages (cluster 1) comprises nearly all EXDO I proteins of validated function. Ten subclusters (A–J) grouping proteins associated with different substrate specificities can be differentiated. Subcluster 1A comprises enzymes experimentally validated as C23O. Interestingly, there is a high redundancy in genomes, as the 28 identified genes are observed in only 18 strains. Out of these, 13 strains belong to the b-proteobacteria and C230 is mainly observed in Burkholderia, Cupriavidus, and Ralstonia genomes. This contrasts previous reports on C23Os, which were predominantly characterized from Pseudomonas strains (Eltis and Bolin, 1996). However, in none of the sequenced Pseudomonas a homologous gene is observed. It has, however, to be noted that most of such genes have previously been reported on plasmids rather than in the chromosome of the strains, such as the case for P. putida KT2440 where the IncP-9 TOL plasmid pWW0 is present (Williams and Murray, 1974), but not included in the same genome project. It is also interesting to note that the Actinobacterium R. jostii RHA1 has a predicted C23O of this kind. Subcluster 1B groups putative homoprotocatechuate 2,3-dioxygenases of the actinobacterial lineage. As expected from literature, the respective encoding genes are present in Actinobacteria (Vetting et al., 2004), and observed in 5 out of 53 genomes. They are absent from any band g-proteobacterial genomes, but surprisingly most abundant in a-proteobacterial genomes

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

. Figure 3 Evolutionary relationships among type I extradiol dioxygenases (EXDO I). Subcluster 1A comprises catechol 2,3-dioxygenases, subcluster 1B putative homoprotocatechuate 2,3-dioxygenases, subcluster 1C proteins related to BphC of Bacillus sp. JF8, subcluster 1D proteins related to NahC of Bacillus sp. JF8, subcluster 1G proteins related to DntD of Burkholderia sp. DNT or BphC3 and BphC4 of R. jostii RHA1, subcluster 1H proteins similar to those capable to cleave 2,3-dihydroxy-p-cumate, subcluster 1I proteins related to those involved in diterpenoid degradation, and subcluster 1J enzymes with similarities to those being active mainly against bicyclic and higher condensed dihydroxylated aromatics. Subcluster 2B comprises so-called one-domain extradiol dioxygenases and cluster 3 proteins related to LinE chlorobenzoquinol 1,2-dioxygenases and PcpA 2,6-dichlorobenzoquinol 1,2-dioxygenases. However, the function of the majority of enzymes of cluster 3 as well as of enzymes of subclusters 1E, 1F, and 2A remains to be elucidated.

(16 genomes), specifically in Bradyrhizobiaceae and Rhodobacteraceae, even though proteobacterial homoprotocatechuate 2,3-dioxygenases are generally assumed to be members of the LigB family (see below) (Roper and Cooper, 1990). It is also interesting to note that such genes were found also outside the Actinobacteria and Proteobacteria, and are present in both sequenced Deinococcus and in both sequenced Thermus strains as well as in three Bacillaceae.

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Subcluster 1C groups proteins related to BphC of Bacillus sp. JF8 involved in biphenyl degradation by this strain (Hatta et al., 2003). Related proteins are not encoded in any of the sequenced Bacilli, but astonishingly in all four genomes available of Chloroflexaceae strains and in a few actinobacterial species, including one protein of R. jostii RHA1, however, not having a taxonomically linked distribution in lower levels. Similarly, proteins related to NahC 1,2dihydroxynaphthalene dioxygenase of Bacillus sp. JF8 (Miyazawa et al., 2004) (subcluster 1D) are not observed in any Bacillus species, but encoded in four a-proteobacterial genomes. Also the three subcluster 1E proteins, where no closely related proteins have been characterized so far, are encoded in two a-proteobacterial genomes. Subcluster 1F proteins are encoded by all 34 genomes available of Burkholderia and various other proteobacterial genomes, however, their actual function still remains to be elucidated. Subcluster 1G comprises proteins such as DntD of Burkholderia sp. DNT responsible for meta-cleavage of trihydroxytoluene, which is also active on catechol (Haigler et al., 1999) but includes as well various proteins of proven activity against 2,3-dihydroxybiphenyl such as BphC3 and BphC4 of R. jostii RHA1, both being reported as being practically inactive with catechol (Sakai et al., 2002). Similar proteins are mainly observed in genomes of Actinobacteria, with R. jostii RHA1 harboring three of such genes, and a- and b-Proteobacteria. Proteins similar to those capable to cleave 2,3-dihydroxy-p-cumate (subcluster 1H) are only found in four genomes including P. putida F1 reported to exhibit such activity (Eaton, 1996) and B. xenovorans LB400, indicating that it is not a widespread activity. Similarly, proteins related to those involved in diterpenoid degradation (subcluster 1I) (Martin and Mohn, 2000), are not common in the genomes analyzed, showing only hits in Caulobacter sp. K31 and the already described activity of B. xenovorans LB400 (Smith et al., 2007). Subcluster 1J comprises a variety of enzymes with similarities to members of subfamilies I.4, I.5, and I.3.E being active mainly against bicyclic and higher condensed dihydroxylated aromatics (Eltis and Bolin, 1996). An overall of 68 such proteins could be observed to be encoded in thus far sequenced genomes. Respective genes are observed in 11 of 17 Mycobacterial genomes, which is not astonishing, as various sequenced Mycobacteria were selected for their capability to mineralize polycyclic aromatics. They are also observed in all three Nocardiaceae genomes, with R. jostii RHA1 harboring six such genes. In addition, eight a-, eight b-, and five g-proteobacterial strains harbor such enzyme. Out of the Pseudomonas, it was observed only in the P. putida F1 genome (Zylstra et al., 1988). The majority of the approximately 100 protein sequences conforming cluster 2 contain the Prosite PS00082 extradiol ring-cleavage dioxygenase signature described above. Subcluster 2B comprises BphC6 of R. jostii RHA1 (ABO34703) and other previously characterized so-called one-domain extradiol dioxygenases such as BphC2 and BphC3 from R. globerulus P6 with reported activity against 2,3-dihydroxybiphenyl (Asturias and Timmis, 1993) (subfamily I.1 as defined by Eltis and Bolin (Eltis and Bolin, 1996)). However, besides BphC6 of strain RHA1, no further enzyme of this type was found to be encoded in the genomes analyzed, and proteins with similarity to subcluster 2A proteins have not yet been functionally characterized. Ring-cleavage dioxygenases involved in the turnover of (chloro)benzoquinols and (chloro)hydroxybenzoquinols have been identified from various microorganisms degrading g-hexachlorocyclohexane or chlorophenols, and comprise LinE chlorobenzoquinol/benzoquinol 1,2-dioxygenases, which preferentially cleaves aromatic rings with two hydroxyl groups at para positions (Miyauchi et al., 1999) and PcpA 2,6-dichlorobenzoquinol 1,2-dioxygenases (Xu et al., 1999). These proteins are comprised in cluster 3, and are the only validated extradiol dioxygenases observed in this cluster. Compared to cluster 1, cluster 3 is so divergent that even

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

the Superfam HMM system recognizes the validated LinE/PcpA sequences as part of the Glyoxalase/bleomycin resistance protein/dioxygenase superfamily but belonging to the family of Glyoxalase I (lactoylglutathione lyase). Only the genomes of Cupriavidus necator H16 and JMP134 contain sequences that may have encode chlorobenzoquinol dioxygenases. It should be noted that one of the sequences of C. necator JMP134 is clustered with a gene similar to the one described from P. putida HS12 encoding nitrobenzene nitroreductase, which is also clustered with a putative benzoquinol extradiol dioxygenase (Park and Kim, 2000).

4.3

Lig B Superfamily

A second family of extradiol dioxygenases is the so-called LigB family (Sugimoto et al., 1999). LigB type extradiol dioxygenases are well established as being responsible for the degradation of protocatechuate via the protocatechuate 4,5-dioxygenase pathway. Protocatechuate dioxygenases are composed of two distinct subunits, with the active site being located in the b-subunit. Also, proteobacterial homoprotocatechuate 2,3-dioxygenases as the one described in Escherichia coli (Roper and Cooper, 1990) belong to the type II or LigB superfamily of extradiol dioxygenases whereas actinobacterial homoprotocatechuate 2,3-dioxygenases are supposed to belong to the EXDO I (Vetting et al., 2004). A further well-documented group of LigB-type extradiol dioxygenases are the 2,3-dihydroxyphenylpropionate 1,2-dioxygenases which, like LigB-type homoprotocatechuate dioxygenases, consist only of one type of subunit (Diaz et al., 2001). Recent analyses have revealed various other substrates that are cleaved by LigB-type extradiol dioxygenases. Aminophenol 1,6-dioxygenases (> Fig. 1) are, like protocatechuate 4,5-dioxygenases, composed of two distinct subunits, with the b-subunits containing the active site (Takenaka et al., 1997). Gallate dioxygenases have so far been described in S. paucimobilis SYK-6 (Kasai et al., 2005) and P. putida KT2440 (Nogales et al., 2005), and are specific for this substrate and do not transform protocatechuate, whereas gallate transformation by protocatechuate 4,5-dioxygenases has been reported. Both gallate dioxygenases have sizes significantly larger than those of the b-subunits of protocatechuate dioxygenases. Analysis of the primary structure revealed that the N-terminal regions showed a significant amino acid sequence identity with the b-subunit of protocatechuate 4,5-dioxygenases, whereas the C-terminal region has similarity to the corresponding small a-subunit (Nogales et al., 2005). It was therefore suggested that gallate dioxygenases are two-domain proteins that have evolved from the fusion of large and small subunits. Additional LigB-type enzymes have been described to be involved in the degradation of methylgallate (Kasai et al., 2004) or of bi- and polycyclic aromatics (Laurie and Lloyd-Jones, 1999). Phylogenetic analysis of the deduced protein sequences of LigB-type proteins encoded in the genomes of bacteria sequenced so far allowed the identification of six clusters (> Fig. 4). Cluster 1 comprises three subclusters, which contain protocatechuate 4,5-dioxygenase b-subunits (> Fig. 4, cluster 1A), gallate dioxygenases (cluster 1B), and a group of related proteins where no member has been characterized thus far (cluster 1C). Respective genes were nearly exclusively observed in a-, b-, and g-Proteobacteria and only 1 of the 53 analyzed actinobacterial genomes (Arthrobacter sp. FB24) has a protocatechuate 4,5-dioxygenase encoding gene. Protocatechuate 4,5-dioxygenases are predominantly observed in Comamonadaceae and Bradyrhizobiaceae, specifically Bradyrhizobium and Rhodopseudomonas strains and are mainly composed of two distinct subunits as evidenced by two subsequent genes encoding the respective subunits. However, putative gene fusions are observed in Arthrobacter and

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Phylogenomics of Aerobic Bacterial Degradation of Aromatics

. Figure 4 Evolutionary relationships among LigB-type dioxygenases. Subcluster 1A comprises protocatechuate 4,5-dioxygenase b-subunits, subcluster 1B gallate dioxygenases, cluster 2, enzymes most closely related to PhnC of Burkholderia sp. strain RP007 or CarBb of P. resinovorans CA10, cluster 3 enzymes related to DesZ of Sphingomonas paucimobilis SYK-6, cluster 4 2,3-dihydroxyphenylpropionates 1,2-dioxygenases, cluster 5 the b- and a-subunits (clusters 5A and B, respectively) of 2-aminophenol 1,6-dioxygenases, and cluster 6 homoprotocatechuate 2,3-dioxygenases. The function of enzymes of subcluster 1C remains to be elucidated.

Verminephrobacter. Even though one of the two gallate dioxygenases characterized so far was reported in a Sphingomonas strain (Kasai et al., 2005), gallate dioxygenase encoding genes are not observed in any of the 112 sequenced a-Proteobacteria and are thus not a dominant trait in this group. In contrast, gallate dioxygenases are obviously encoded in the genomes of three of four sequenced P. putida strains. The supposed gallate dioxygenases are mainly fusions of a- and b-subunits, like in P. putida KT2440 (Nogales et al., 2005), however, seem to consist of separate subunits in Xanthomonas and Chromohalobacter. Dioxygenases belonging to the third subcluster are usually composed of a- and b-subunits, and are in 10 out of 12 cases encoded in genomes, which also encode a protocatechuate 4,5-dioxygenase pathway. A second cluster (cluster 2, > Fig. 4) comprises enzymes most closely related to those involved in bi- and polycyclic aromatic degradation such as PhnC involved in the degradation

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

of polycyclic aromatics by Burkholderia sp. strain RP007 (Laurie and Lloyd-Jones, 1999), CarBb involved in the degradation of carbazol by P. resinovorans CA10 (Sato et al., 1997a), or BphC6 involved in the degradation of fluorene by Rhodococcus rhodochrous K37 (Taguchi et al., 2004). However, no clear association with a capability to degrade such compounds was evident, and the respective enzymes are spread among very different groups of Actinobacteria and Proteobacteria. The corresponding genes are absent from strains selected for genome sequencing due to their exceptional capability to degrade aromatics such as M. vanbaalenii Pyr, M. gilvium PYR-GCK, R. jostii RHA1, or B. xenovorans LB400. Cluster 3 comprises enzymes related to DesZ methylgallate dioxygenase of Sphingomonas paucimobilis SYK-6, where 7 out of 11 proteins are observed in Mycobacterium strains, however, their function remains to be elucidated. A fourth cluster obviously comprises 2,3-dihydroxyphenylpropionate 1,2-dioxygenases. The respective enzymes are most dominantly observed to be encoded in the genomes of Enterobacteriaceae, and specifically observed in 13 out of 18 E. coli strains sequenced and in Shigella sonnei. Interestingly, related enzymes are also observed to be encoded by 9 out 17 Mycobacterial genomes. Their function, however, remains to be proven. A fifth cluster comprises 2-aminophenol 1,6-dioxygenases (> Fig. 4, clusters 5A and B comprising the b- and a-subunits, respectively). Only two of these enzymes are observed to be encoded by previously sequenced genomes, i.e., B. xenovorans LB400 and P. putida W619, indicating such pathways to be present only in very few specialized bacteria. In contrast, homoprotocatechuate 2,3-dioxygenases (cluster 6) are observed to be widespread, and in contrast to previous assumptions that LigB-type homoprotocatechuate 2,3-dioxygenases were restricted to proteobacteria, homologues are also observed in two Actinobacteria, and the genomic context suggest that those enzymes actually are part of a functional homoprotocatechuate pathway. A homologue is also observed in Bacillus licheniformis.

4.4

Cupin Dioxygenases

Several extradiol dioxygenases of aromatic degradation pathways have been described to belong to the cupin superfamily (Dunwell et al., 2000) sharing a common architecture and including key enzymes such as gentisate 1,2-dioxygenase (involved in the degradation of salicylate or 3-hydroxybenzoate, > Fig. 1), homogentisate 1,2-dioxygenase (involved in the degradation of phenylalanine and tyrosine) (Arias-Barrau et al., 2004) and 3-hydroxyanthranilate 3,4-dioxygenase (involved in tryptophan degradation) (Kurnasov et al., 2003; Muraki et al., 2003). The phylogenomic analysis of this type of dioxygenases in the genomes of bacteria sequenced so far shows that homogentisate dioxygenase is the enzyme with the broadest distribution in bacterial families. This may be explained by the key role in the degradation of the aromatic amino acids phenylalanine and tyrosine in several organisms, including eukaryotes. Putative genes encoding this enzyme are strongly represented in Proteobacteria, being identified in 10 out of 19 a-, 5 out of 10 b-, 16 out of 29 g-, and 4 out of 11 d-proteobacterial families, although they were absent in e-proteobacteria. In the families Bradyrhizobiaceae, Rhizobiaceae, Alcaligenaceae, Burkholderiaceae, Shewanellaceae, Legionellaceae, Pseudomonadaceae, and Vibrionaceae, a respective gene can be observed in nearly all genomes sequenced. Homogentisate 1,2-dioxygenase was the unique aromatic ring-cleavage enzyme found in sequenced representatives of the families Hyphomonadaceae, Neisseriaceae, Aeromonadaceae, Idiomarinaceae, Moritellaceae, Chromatiaceae, Legionellaceae, Hahellaceae, Bdellovibrionaceae,

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Cystobacteraceae, and Nannocystaceae. In addition, genes putatively encoding homogentisate 1,2-dioxygenase are also found in members of the non-proteobacterial orders Actinomycetales, Flavobacteriales, Sphingobacteriales, and Bacillales. Gentisate 1,2-dioxygenase is the ring-cleavage enzyme involved in catabolism of salicylate and 3-hydroxybenzoate, among other aromatics (> Fig. 1). In comparison to homogentisate 1,2-dioxygenases, gentisate 1,2-dioxygenases show a narrow distribution in bacterial families of proteobacteria being identified only in six a-, three b-, and three g-proteobacterial families and being absent from d- and e-proteobacteria. The number of members with putative gentisate 1,2-dioxygenase genes inside the 12 proteobacterial families owing this enzyme is also significantly lower than the percentage of homogentisate 1,2-dioxygenase carrying members. Inside the Comamonadaceae however, six out of eight members harbor a gentisate 1,2-dioxygenase, but only one a homogentisate dioxygenase. Similarly, homogentisate dioxygenases are absent from the genomes of Enterobacteriaceae, although Salmonella, Serratia, and some E. coli strains are endowed with a gentisate dioxygenase. In addition to Proteobacteria, gentisate 1,2-dioxygenase genes can be found in Corynebacteriaceae, Micrococcaceae, Mycobacteriaceae, Nocardiaceae, and Bacillaceae. 3-Hydroxyanthranilate 3,4-dioxygenase catalyzes the conversion of 3-hydroxyanthranilate to 2-amino-3-carboxymuconic semialdehyde during tryptophan degradation via the kynurenine pathway. This extradiol dioxygenase is the cupin-type dioxygenase with the narrowest distribution since it is only found and with a low representativity in Brucellaceae, Rhodobacteraceae, Sphingomonadaceae, Burkholderiaceae, Shewanellaceae, Xanthomonadaceae, and Myxococcaceae in Proteobacteria and in Flavobacteriaceae, Flexibacteraceae, and Bacillaceae in non-proteobacterial families.

4.5

Other Extradiol Dioxygenases

Recently, a novel Fe2+-dependent dioxygenase, benzoquinol 1,2-dioxygenase, which is a a2b2 heterotetramer where the a- and b-subunits displayed no significant sequence identity with other dioxygenases and which catalyzes the ring fission of a wide range of benzoquinols to the corresponding 4-hydroxymuconic semialdehydes, has been described in P. fluorescens ACB (Moonen et al., 2008). Putative genes encoding both subunits of benzoquinol 1,2-dioxygenase show a highly narrow distribution since they are almost exclusively found in Burkholderia with the exceptions of P. luminescens subsp. laumondii TTO1 and P. aeruginosa PA7 strains, in spite to be originally identified in a 4-hydroxyacetophenone-degrading P. fluorescens strain (Moonen et al., 2008). The origin of this type of dioxygenase remains to be clarified.

4.6

Diiron Oxygenases

Soluble diiron oxygenases comprise an evolutionary-related family of enzymes capable to monooxygenate benzene/toluene to phenol/methylphenol and phenols to catechols (Leahy et al., 2003). Sequence comparisons of the respective a-subunits with the PaaA oxygenase subunit of phenylacetyl-CoA oxygenase and the BoxB oxygenase of benzoyl-CoA oxygenase strongly suggest that also these enzymes belong to the family of soluble diiron oxygenases. Benzene/toluene monooxygenases and phenol monooxygenases of the soluble diiron oxygenase family are enzyme complexes including an electron transport system comprising

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

a reductase (and, in some cases, a ferredoxin), a catalytic effector and a terminal heteromultimeric oxygenase composed by a, b, and g subunits whose a-subunits are assumed to be the site of substrate hydroxylation (Leahy et al., 2003). According to the presence of genes putatively coding for a subunit, benzene/toluene multicomponent monooxygenase are found almost exclusively in b-Proteobacteria, including Burkholderia, Cupriavidus, Ralstonia, Methylibium, and Dechloromonas strains with the only exceptions of Bradyrhizobium sp. BTAi1 and Frankia sp. CcI3. In the b-proteobacterial strains, the benzene/toluene multicomponent monooxygenase are associated with a phenol/methylphenol multicomponent monooxygenase. On the other hand, the phenol/methylphenol multicomponent monooxygenases showed a slightly broader distribution since in addition to the above mentioned strains, such genes are also identified in Acidovorax and Verminephrobacter strains and even in g-proteobacterial families such as Alteromonadaceae and Pseudomonadaceae. In contrast to the limited distribution of the above described multicomponent monooxygenases, multicomponent phenylacetyl-CoA oxygenases are broadly distributed in Proteobacteria being identified in 6 out of 19 a-, 5 out of 10 b-, and 8 out of 29 g-proteobacterial families. They are, however, absent from d- and e-proteobacteria. The families Rhodobacteraceae, Bradyrhizobiaceae, Alcaligenaceae, Burkholderiaceae, Rhodocyclaceae, Enterobacteriaceae, and Pseudomonadaceae include a significant number of strains with such genes. Several representatives are also found in non-proteobacterial families, predominantly Actinobacteria such as Streptomycetaceae, Pseudonocardiaceae, Nocardiaceae, Micrococcaceae, Corynebacteriaceae, Brevibacteriaceae, and Acidothermaceae, and also in Flavobacteriaceae and Bacillaceae families. Benzoyl-CoA oxygenase encoding genes are exclusively found in some families of the a- and b-proteobacteria: Bradyrhizobiaceae, Rhodospirillaceae, Comamonadaceae, Burkholderiaceae, and Rhodocyclaceae, and predominantly in the last two families in which the pathway was also originally described (Denef et al., 2004; Zaar et al., 2004).

4.7

Flavoprotein Monooxygenases

Flavoprotein monooxygenases are involved in a wide variety of biological processes including biosynthesis of antibiotics and siderophores or biodegradation of aromatics. They have been classified according to sequence and structural data in six classes (van Berkel et al., 2006), with classes A, D, and F being of special importance for aromatic degradation. Class A enzymes are considered to be widely distributed in different bacterial taxa and typically ortho- or parahydroxylate aromatic compounds that contain an activating hydroxyl- or amino-group (van Berkel et al., 2006). In fact, it is interesting to note that according to genome annotations, a huge set of bacteria contain enzymes capable of 4-hydroxybenzoate 3-hydroxylation, salicylate 1-hydroxylation or 2,4-dichlorophenol 6-hydroxylation. Regarding the fact that the capability to mineralize chloroaromatics is not widespread in bacteria and chlorocatechol genes, usually necessary to achieve mineralization of chloroaromatics are, among the sequenced genomes only observed in the two bacteria well studied for such capability, i.e., B. xenovorans LB400 (Chain et al., 2006) and C. necator JMP134 (Pe´rez-Pantoja et al., 2008), the annotated widespread of enzymes involved in dichlorophenol degradation is astonishing. A phylogenetic analysis of proteins related to enzymes of class A flavoproteins using proteins of documented function (salicylate 1-hydroxylases, 3-hydroxybenzoate 4-hydroxylases, 2-aminobenzoyl-CoA monooxygenases/reductases, 4-hydroxybenzoate 3-hydroxylases, among others) as seeds show that these oxygenases can be grouped into six distinct protein clusters (enzymes related to

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Phylogenomics of Aerobic Bacterial Degradation of Aromatics

UbiH involved in ubiquinone biosynthesis will not be discussed here). Only one of these clusters comprises enzymes, which, based on characterized representatives, can be assumed to catalyze a single defined activity, i.e., the 3-hydroxylation of 4-hydroxybenzoate. As with the majority of aromatic degradative properties, the respective enzymes are predominantly observed in Actinobacteria and Proteobacteria. However, they are also observed in one of two Acidobacteria, in Pedobacter of the Bacteroidetes, in one Deinococcus and in 1 of 28 Bacillaceae. No other monocomponent flavoprotein monooxygenases discussed in this section are observed in these orders. Among the Actinobacteria, 4-hydroxybenzoate 3-hydroxylases are observed in roughly one third of the families, including Arthrobacter and Streptomyces, but interestingly were absent from any of the 17 Mycobacterium analyzed. It is a dominant trait in a-Proteobacteria, specifically in Bradyrhizobiaceae and Rhodobacteraceae. Also among b-Proteobacteria, all 34 Burkholderia, three Cupriavidus, four Ralstonia, and six out of eight Comamonadaceae are endowed with such capability. In contrast, such activity is rare in g-Proteobacteria with the exception of Pseudomonadaceae, where 17 out of 18 strains (exception again P. mendocina) have a 4-hydroxybenzoate 3-hydroxylase. Similarly, such activity is spread among Acinetobacter and Xanthomonas strains. Among the Enterobacteriaceae, only Klebsiella pneumoniae and Serratia proteomaculans have a 4-hydroxybenzoate 3-hydroxylase. Also the aminobenzoyl-CoA pathway (Altenschmidt and Fuchs, 1992) seems to be strongly represented among the thus far sequenced bacteria. In a phylogenetic analysis, the aminobenzoyl-CoA oxygenases seem to be related to salicylyl-CoA 5-hydroxylase from Streptomyces sp. WA46 (Ishiyama et al., 2004) channeling salicylate to gentisate. However, in contrast to the organization in strain WA46 where the oxygenase encoding gene is clustered with a gentisate dioxygenase, function as a salicylyl-CoA 5-hydroxylase can be suggested only in a few cases, such as in S. wittichii RW1, since a gentisate pathway is absent from the genomes of various strains including the two Streptomyces strains sequenced. Overall, homologues to aminobenzoyl-CoA oxygenases are observed in 44 genomes comprising Actinobacteria (five genomes) such as Streptomyces or Saccharopolyspora erythraea NRRL 2338. In Proteobacteria this pathway is absent in g-Proteobacteria, but it is observed in Plesiocystis pacifica SIR-1 (a d-proteobacterium). The pathway is abundant in b-Proteobacteria such as Azoarcus strains, where this metabolic route was initially established (Altenschmidt and Fuchs, 1992), but also in Comamonadaceae (six of eight genomes), Ralstonia (all four genomes), Cupriavidus (all three genomes), and a-proteobacteria such as Bradyrhizobium strains (all three genomes) or Rhodobacteraceae (11 of 24 genomes). A large number of genes in bacterial genomes (nearly 100) are annotated as encoding salicylate 1-hydroxylases. However, a phylogenetic analysis taking into account validated salicylate 1-hydroxylases, identified only two of such proteins (amino acid sequence identity >40% to validated NahG proteins [Yen and Gunsalus, 1982]) encoded in the genome of A. baylyi ADP1 (as previously described [Jones et al., 2000]) and P. putida GB-1 (see > Fig. 5, cluster 1). Also enzymes related to NahW, a second evolutionary lineage of salicylate 1-hydroxylases (Bosch et al., 1999b) are scarce and only seven homologues (four of them encoded by Burkholderia genomes) are identified (sequence identity >35%) (see > Fig. 5, cluster 5). In contrast, various enzymes (observed in 22 genomes) clustered with enzymes of proven function as 3-hydroxybenzoate 6-hydroxylases (> Fig. 5, cluster 10) and were observed, among others, in three Corynebacteria, two Arthrobacter, seven Burkholderiaceae, and three Comamonadaceae strains. Other enzymes annotated as salicylate hydroxylases (16) show high similarity (>60% identity) and cluster together with 6-hydroxynicotinate 3-monooxygenase of P. fluorescens TN5 (Nakano et al., 1999) such that their function as salicylate hydroxylases is

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

. Figure 5 Evolutionary relationships among proteins related to NahG, or NahW-type salicylate 1-hydroxylases and 3-hydroxybenzoate 6-hydroxylases. Clusters 1 and 5 comprise salicylate 1-hydroxylases related to NahG or NahW salicylate 1-hydroxylases, cluster 10 3-hydroxybenzoate 6-hydroxylases, and cluster 2 enzymes related to 6-hydroxynicotinate 3-monooxygenase of Pseudomonas fluorescens TN5. The function of enzymes of other clusters remains to be elucidated.

questionable (> Fig. 5, cluster 2). The same holds true for a further more than 100 additional sequences, out of which 69 (> Fig. 5, cluster 6–9) are, among enzymes with validated function, phylogenetically most closely related to 3-hydroxybenzoate 6-hydroxylases. However their genomic contexts indicate different functions. A similar situation holds for enzymes annotated as 3-hydroxyphenylpropionate monooxygenases. An overall of 24 proteins showed significant similarity (>40% identity) with respective validated enzymes and, in phylogenetic analysis, clustered together in one evolutionary branch. These enzymes are predominantly observed in Mycobacterium (seven genomes) and Enterobacteriaceae (mainly E. coli, 11 genomes, but also in K. pneumoniae and S. sonnei), as well as in B. vietnamiensis, B. xenovorans, C. necator JMP134, and P. putida W619. Other enzymes annotated as 3-hydroxyphenylpropionate monooxygenases show significant similarity to either resorcinol monooxygenase of C. glutamicum (Huang et al., 2006) or to GdmM involved in formation of the geldanamycin benzoquinoid system by S. hygroscopicus

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AM 3672 (Rascher et al., 2005) and are thus highly improbable to function as 3-hydroxyphenylpropionate monooxygenase. A 3-hydroxyphenylacetate 6-hydroxylase forming homogentisate has been recently described in P. putida U being composed of the hydroxylase and a small coupling protein, constituting a novel type of two-component hydroxylase, distinct from the classical twocomponent flavoprotein monooxygenases (Arias-Barrau et al., 2005). Seventeen homologues (>40% sequence identity, clustering on the same phylogenetic branch) are observed in 16 of the so far sequenced genomes and usually two subsequent genes encoding for the coupling protein and the monooxygenase can be identified. Interestingly, in contrast to the first and thus far only observation in Pseudomonas, such genes are absent from all 17 sequenced Pseudomonas strains and all other g-proteobacterial genomes but frequently found in Burkholderia (5 of 34 genomes), Cupriavidus (two of three genomes), and Comamonadaceae (four out of eight genomes). Also, various flavoprotein monooxygenases are annotated as 2,4-dichlorophenol hydroxylases. However, enzymes related to valid 2,4-dichlorophenol hydroxylases (>40% sequence identity) also comprise phenol hydroxylases such as PheA from Pseudomonas sp. strain EST1001, which transforms phenol and 3-methylphenol, but not 2,4-dichlorophenol (Nurk et al., 1991), ChqA chlorobenzoquinol monooxygenase of Pimelobacter simplex (AY822041), HpbA 2-hydroxybiphenyl-3-monooxygenase from P. azelaica HBP1, which is capable of oxidizing various 2-substituted phenols, but not phenol (Suske et al., 1997), OhpB 3-(2-hydroxyphenyl)propionic acid monooxygenase from R. aetherivorans I24 (DQ677338) and MhqA methylbenzoquinol monooxygenase from Burkholderia NF100 (Tago et al., 2005). Thus, enzymes of this group typically share the capability to transform 2-substituted phenols, but are obviously recruited for different metabolic routes and involve pathways where the ring-cleavage substrate is a dihydroxylated compound, but also routes where the ring-cleavage substrate is trihydroxylated. The function of these proteins, therefore, cannot be deduced from similarity measures or from phylogenetic analysis. An overall of 18 proteins can be identified as belonging to this cluster, and beside the two characterized 2,4-dichlorophenol hydroxylases from C. necator JMP134 only two genomes (Rhizobium leguminosarum and Bradyrhizobium sp. ORS278) comprise proteins clustering with 2,4-dichlorophenol hydroxylases. However, the genetic environment of the encoding genes does not give a direct support for such a function. Further proteins of this cluster are observed to be scattered among Actinobacteria and Proteobacteria with R. jostii RHA1 encoding for three of such proteins. Interestingly, a distinct group of flavoprotein monooxygenases exhibiting approximately 30% of sequence identity to the above described monooxygenases is also typically annotated as phenol hydroxylases. This annotation seems to be due to some similarity to the phenol hydroxylase (30–35% identity) of Trichosporon cutaneum (Enroth et al., 1994), however, phylogenetic analysis shows that a set of 29 proteins (typically with identities >50%) is most closely related to proteins of validated function as 3-hydroxybenzoate 4-hydroxylases, previously assumed to be restricted to Comamonas strains (Hiromoto et al., 2006). In fact, inside the b-proteobacteria such genes are only observed in C. testosteroni and B. phymatum, however, also three g-Proteobacteria harbor such gene, and 3-hydroxybenzoate-4-hydroxylases seem to be frequently encoded in the genome of a-Proteobacteria (12 genomes), specifically in Bradyrhizobium strains (all three genomes) and Rhodobacteraceae (6 out of 24 genomes). Also seven Actinobacteria seem to harbor such activity (among them two Corynebacterium species and both sequenced Arthrobacter strains), indicating this activity to be more widespread than previously thought.

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39

Nearly 20 enzymes were annotated as pentachlorophenol monooxygenases, an activity previously reported, for example, in Sphingobium chlorophenolicum (Cai and Xun, 2002). However, none of these proteins showed sequence identities >35% to validated PcpB proteins, and only a group of enzymes typically encoded in Burkholderia genomes could be shown to be evolutionary related, however, their function as PCP monooxygenases seems highly improbable. Styrene monooxygenases (StyA) have been identified in various Pseudomonas strains (Beltrametti et al., 1997), and were classified as Class E flavoprotein monooxygenases, however, they are evolutionary related to the Class A flavoprotein monooxygenases (van Berkel et al., 2006). Interestingly, none of the sequenced Pseudomonas strains harbor such a gene. Eight phylogenetically related proteins are observed in genome sequencing projects, however their function as such monooxygenases remains speculative. Two-component aromatic hydroxylases such as 4-hydroxyphenylacetate 3-hydroxylases from E. coli (Diaz et al., 2001) consisting of an oxidoreductase and an oxygenase were classified as type D flavoprotein monooxygenases (Ballou et al., 2005) and have no structural or sequence similarities to the single-component enzymes described above. Iterative Psi-blast searches identified nearly 100 of such enzymes putatively involved in aromatic metabolism to be encoded in sequenced genomes and phylogenetic analysis indicated the presence of eight evolutionary lines (see > Fig. 6). Two of the branches contain the proteobacterial (> Fig. 6, cluster 1) and non-proteobacterial (> Fig. 6, cluster 7) 4-hydroxyphenylacetate 3-hydroxylases with an identity of members of the different cluster of approximately 30%. Proteins located on the same phylogenetic branch as validated 4-hydroxyphenylacetate 3-hydroxylases from Thermus or Geobacillus (Hawumba et al., 2007; Kim et al., 2007) are observed in only three Actinobacteria, but in both sequenced Deinococci and in both Thermus strains. It is also a dominant trait in Bacillaceae (13 out of 28 genomes). Among the Proteobacteria, 4-hydroxyphenylacetate 3-hydroxylation by enzymes of this cluster is a trait nearly exclusively observed in g-proteobacteria, predominantly in Enterobacteriaceae (19 out of 61 genomes) and Pseudomonas (5 out of 18 genomes), and outside of this group only in two a-proteobacteria. The cluster of proteins most closely related to these proteobacterial 4-hydroxyphenylacetate 3-hydroxylases (50–60% identity) comprises those with high similarity to phenol hydroxylase PheA of Geobacillus thermoleovorans (Duffner and Muller, 1998), R. erythropolis (CAJ01325), 4-nitrophenol hydroxylase of Rhodococcus sp. PN1 (Takeo et al., 2003), an enzyme which also acts as a phenol hydroxylase, and 4-coumarate 3-hydroxylase of Saccarothrix espanaensis involved in the formation of caffeic acid (Takeo et al., 2003) (see > Fig. 6, cluster 2). Interestingly, respective genes are practically absent from proteobacteria and only observed in Photorhabdus and Saggitula, but observed in one of the two Thermus strains sequenced, in all Chloroflexaceae and in some Actinobacteria such as R. jostii RHA1, which harbors four homologues. A further group of proteins show similarity to PvcC, previously assumed to be involved in pyoverdin synthesis, but recently shown to be involved in the formation of pseudoverdine and paerucumarin by P. aeruginosa (Takeo et al., 2003) (> Fig. 6, cluster 3). Interestingly, respective genes and gene clusters are exclusively observed in P. aeruginosa, B. mallei, B. pseudomallei, and B. thailandensis. A further cluster of six proteins, also typically annotated as 4-hydroxyphenylacetate 3-hydroxylases is related to TcpA 2,4,6-trichlorophenol monooxygenases of C. necator

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. Figure 6 Evolutionary relationships among the large subunits of two-component flavoprotein monooxygenases related to 4-hydroxyphenylacetate 3-hydroxylase from Escherichia coli. Clusters 1 and 7 comprise 4-hydroxyphenylacetate 3-hydroxylases of proteobacteria and non-proteobacteria, cluster 2 proteins related with PheA phenol hydroxylase of Geobacillus thermoleovorans, and cluster 3 proteins with similarity to PvcC of P. aeruginosa (Takeo et al., 2003) (> Fig. 6, cluster 3). The function of enzymes of other clusters remains to be elucidated.

JMP134 (Sanchez and Gonzalez, 2007), however, the function of these proteins also remains to be elucidated (> Fig. 6, cluster 8). A different type of two-component aromatic hydroxylases consisting also of a reductase and an oxygenase has been described recently (Thotsaporn et al., 2004). This type has been also classified as type D flavoprotein monooxygenases (Ballou et al., 2005) but it is able to use FMN, FAD, and riboflavin for hydroxylation in contrast to HpaB, PheA, and TcpA, which specifically uses only reduced FAD (Thotsaporn et al., 2004). The best studied representative of this group is 4-hydroxyphenylacetate 3-hydroxylase from A. baumannii but it shows very low identity with the 4-hydroxyphenylacetate 3-hydroxylases described previously in E. coli, P. aeruginosa, or T. thermophilum (Thotsaporn et al., 2004). Although the different types of 4-hydroxyphenylacetate 3-hydroxylase catalyze the same reaction, they have significant differences in the details of the mechanisms involved (Ballou et al., 2005). Genes putatively coding for enzymes similar to the A. baumannii-type of 4-hydroxyphenylacetate 3-hydroxylase are found in some strains of a- and g-proteobacteria: S. stellata, R. sphaeroides, Marinomonas sp., V. shilonii, V. vulnificus, A.vinelandii, P. entomophila, and one P. putida strain. Additional

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39

enzymes of this kind of two-component aromatic hydroxylases includes naphthoate 2-hydroxylase (NmoAB) described in Burkholderia sp. JT1500 (Deng et al., 2007) with homologous genes in some Bradyrhizobium and Cupriavidus strains and resorcinol hydroxylase from Rhizobium sp. MTP-10005 (GraAD) (Yoshida et al., 2007) with homologous genes in the related strains A. tumefaciens and R. leguminosarum and in the b-proteobacterium Polaromonas sp. JS666.

4.8

Rieske Non-Heme Iron Oxygenases

The so-called Rieske non-heme iron oxygenases are one of the key families of enzymes important for aerobic activation and thus degradation of aromatics such as benzoate, benzene, toluene, phthalate, naphthalene, or biphenyl (> Fig. 1) (Gibson and Parales, 2000). Members of this family also catalyze monooxygenations, such as salicylate 1- or salicylate 5-hydroxylases or demethylations, such as vanillate O-demethylases. They are multicomponent enzyme complexes consisting of a terminal oxygenase component (iron–sulfur protein [ISP]) and electron transport proteins (a ferredoxin and a reductase or a combined ferredoxin-NADHreductase). The catalytic ISPs are usually heteromultimers composed of a large a-subunit containing a Rieske-type [2Fe-2S] cluster, with a mononuclear nonheme iron oxygen activation center, and a substrate-binding site modulating substrate specificity and a small b-subunit, however, some enzymes, such as phthalate 4,5-dioxygenases contain an oxygenase composed only of a-subunits. Phylogenetic analyses of Rieske non-heme iron oxygenases show that sequences obtained in our searches can be grouped into three main divergent clusters or divisions, where only two of them comprise proteins of validated function and are thus discussed here. One of these two divisions comprises the so-called phthalate family including vanillate demethylases (Gibson and Parales, 2000). Four clusters of this division contain oxygenases of proven function to dioxygenate aromatics, i.e., phthalate 4,5-dioxygenases (Nomura et al., 1992), isophthalate dioxygenase (Wang et al., 1995), phenoxybenzoate dioxygenase (Dehmel et al., 1995), and carbazol dioxygenase (Sato et al., 1997b). Genes putatively encoding phthalate 4,5-dioxygenases are nearly exclusively observed in b-proteobacteria (seven genomes) except for an amazing five homologues possibly encoded in the genome of Rhodobacterales bacterium HTCC2654. Similarly, genes putatively encoding isophthalate dioxygenases are predominantly observed in b-proteobacterial genomes (overall in five), but also in one g-proteobacterium and in two a-proteobacteria, among them strain HTCC2654. A similar spread is observed for enzymes related to phenoxybenzoate dioxygenase (observed in seven b-, four a-, and one gProteobacterium). Genes putatively encoding carbazol dioxygenases are not observed in any sequenced genome. Most of the currently characterized Rieske non-heme iron oxygenases are concentrated in a well-defined division (see > Fig. 7). The significant amount of validly described enzymes allows assignment of putative functions to most of the respective enzymes encoded in sequenced genomes. Benzoate dioxygenases (cluster A1) are most widely distributed and can be observed in the genomes of Actinobacteria as well as a-, b-, and g-proteobacteria. Most importantly, such enzymes are observed in 32 out of 34 Burkholderia strains, 14 out of 18 Pseudomonas strains, and 4 out of 17 Mycobacteria. Anthranilate can be transformed either by two-component anthranilate dioxygenases such as the one described from Acinetobacter baylyi ADP1

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. Figure 7 Evolutionary relationships among the a-subunits of Rieske non-heme iron oxygenases excluding phthalate family enzymes. A function can be assigned to proteins of some of the clusters shown as follows: cluster A1, benzoate dioxygenases; cluster A2, two component anthranilate dioxygenases; cluster A3, proteins related with p-cumate dioxygenases; cluster B3, aniline dioxygenases; cluster C1, NidA-type dioxygenases; cluster C2, phthalate 3,4-dioxygenases; cluster C3, proteins related with diterpenoid dioxygenases; cluster C5, NahA-type naphthalene dioxygenases; cluster 6, proteins related with ethylbenzene dioxygenase from R. jostii RHA1; cluster C8, 3-phenylpropionate dioxygenases; cluster C9, benzene/toluene/isopropylbenzene/ biphenyl dioxygenases; cluster E1, salicylate 5-hydroxylases; cluster E2, 2-chlorobenzoate dioxygenases; cluster E3, terephthalate dioxygenases; cluster E4, salicylate 1-hydroxylases; and cluster E5, three component anthranilate dioxygenases. The function of enzymes of other clusters remains to be elucidated.

(Eby et al., 2001) (cluster A2) or by three-component anthranilate dioxygenase as the one from Burkholderia cepacia DBO1 (Chang et al., 2003) (cluster E5). Genome analysis clearly showed that two-component dioxygenases are obviously restricted to g-proteobacteria and are only observed in seven Pseudomonas genomes and, as described, in A. baylyi. In contrast, three-component anthranilate dioxygenases are exclusively observed in Burkholderia genomes and present in 31 out of 34 sequenced strains. Cluster A3 comprises proteins phylogenetically

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39

related with known p-cumate dioxygenases. These sequence relatives are found in five of the sequenced Pseudomonas genomes but also in S. wittichi RW1 and B. xenovorans LB400. Cluster B3 comprises proteins similar to aniline dioxygenases, and similar sequences are found only in Nocardioides sp. JS614 and Bradyrhizobium sp. BTAi1, indicating a very restricted distribution of such activity. Further related sequences, where no specific function can be postulated (clusters B1, B2, and B4) were predominantly observed in Burkholderiaceae. Proteins of cluster C1 exhibit similarity to proteins involved in the degradation of polycylic aromatics by Actinobacteria, exemplified by NidA of M. vanbaalenii PYR-1 (Stingley et al., 2004a) and thus putatively have a function in degradation of polycyclic aromatics. In accordance with this assumption, respective proteins are found to be encoded in the genomes of five environmental Mycobacteria and up to four different such proteins are observed per genome. As NidA-like proteins, also sequences putatively encoding phthalate 3,4-dioxygenases (Stingley et al., 2004b) (cluster C2) are exclusively observed in Actinobacteria, differentiating them from b-proteobacteria which obviously degrade phthalate by phthalate 4,5-dioxygenases. Phthalate 3,4-dioxygenases were observed to be encoded in genomes of Mycobacteria comprising a NidA sequence, but also in M. avium strains, R. jostii RHA1, and Arthrobacter sp. FB24. Group C3 proteins, comprising diterpenoid dioxygenases-like proteins (Martin and Mohn, 1999) are having a very restricted distribution in the genomes available so far, being found only in Caulobacter sp. K31, Sphingomonas sp. SKA58, S. wittichii RW1, and B. xenovorans LB400 genomes (Smith et al., 2007). Naphthalene and phenanthrene dioxygenases related to NahA of P. stutzeri AN10 (Bosch et al., 1999a) have previously been observed in various Pseudomonas, Sphingomonas, Burkholderia, Cycloclasticus, Acidovorax, and Ralstonia isolates. The genomic survey indicates such activities (see cluster C5) not to be widespread and similar sequences are only observed in genomes of N. aromaticivorans DSM 12444, Acidovorax sp. JS42, and P. naphthalenivorans CJ2. Also sequences related to ethylbenzene dioxygenase from strain RHA1 (Iwasaki et al., 2006) (cluster C6) are additionally observed only in of Azotobacter vinelandii AvOP and N. aromaticivorans DSM 12444. Sequences indicating to encode 3-phenylpropionate dioxygenases (cluster C8) are exclusively observed in Enterobacteriaceae, and interestingly observed in all Shigella spp. strains (seven genomes) and 11 of 17 E. coli. Cluster C9 is composed of benzene/toluene/isopropylbenzene/biphenyl dioxygenases (Witzig et al., 2006), enzymes typically involved in the degradation of the respective compounds, where a broad set of both proteobacterial and actinobacterial isolates is available. Respective sequences are only observed in the four genomes of strains previously reported to harbor such activity (P. putida F1, B. xenovorans LB400, P. napthalenivorans CJ2, and R. jostii RHA1). Cluster E comprises enzymes acting on ortho- or para-substituted benzoates and include salicylate 5-hydroxylases (Fuenmayor et al., 1998) (cluster E1), salicylate 1-hydroxylases (Pinyakong et al., 2003) (cluster E4), 2-chlorobenzoate dioxygenases (cluster E2), threecomponent anthranilate dioxygenases (cluster E5, see above), and terephthalate dioxygenases (Sasoh et al., 2006) (cluster E3). Respective sequences are nearly exclusively observed in b-proteobacteria and in Sphingomonads out of the a-proteobacteria and only terephthalate dioxygenases are also observed in Actinobacteria, i.e., R. jostii RHA1 and Arthrobacter aurescens T1, which corresponds with various reports of Rhodococci being capable of degrading terephthalate. Terephthalate dioxygenases are also observed in B. xenovorans LB400 and

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C. testosteroni with members of last mentioned genus also often being implicated in terephthalate degradation (Sasoh et al., 2006). Salicylate 5-hydroxylases were observed in two Cupriavidus strains, both Polaromonas strains, and both R. solanacearum isolates in accordance with such activity being first described from a Ralstonia strain (Fuenmayor et al., 1998). Also S. wittichii RW1 seems to harbor such activity. In contrast, a Rieske-type salicylate 1-hydroxylase was only observed in N. aromaticivorans DSM 12444, also in accordance with the fact that such activities so far have only been described in Sphingomonads. Also putative 2-chlorobenzoate 1,2-dioxygenases are rare and a putative homologue is only observed in the genome of B. xenovorans LB400.

5

Metabolism Diversity

A very exciting question can be addressed based on the phylogenomic analyses carried out here: What is the diversity of catabolic properties within phylogenetic groups? However, before answering such question, a definition about the ‘‘unit of catabolic diversity’’ must first be addressed. The first unit level is pathway diversity. It refers to the presence in one bacterium or bacterial group of different ways to degrade one compound (i.e., intradiol versus extradiol ring cleavage; classical aromatic ring oxidation versus a CoA-dependent pathway, etc.). This level of diversity is the thickest and provides the most powerful versatility because it allows the microorganism to choose among very different ways to metabolize the compound. The second level of ‘‘unit of diversity’’ is the enzymatic diversity. It refers to the same biochemical reaction or catabolic step carried out by completely different enzymes. For example, enzymes belonging to three different families can perform phenol conversion to catechol: single-component flavoprotein monoxygenases, diiron oxygenases, or two-component monooxygenases. This level of catabolic diversity is finer than the previous one, but still significant because it allows for versatility at the biochemical level, i.e., different substrate affinities, different cofactor requirements, inhibitor effects, among others. The third level of catabolic diversity is the genetic diversity, or classical gene redundancy: the same biochemical step may be performed by very similar enzymes encoded by different genes. It is assumed that the main point of diversity here is at the regulatory level. Although a gross measure of catabolic versatility, in the following three sections the pathway diversity will be used as a diversity unit for aromatic catabolism properties of a taxonomic group. This is especially relevant to account for the diversity of central pathways as defined in > Table 2.

5.1

Metabolism by Bacteria Outside the Actinobacterial and Proteobacterial Phyla

When the genome database is searched for the aromatic catabolic pathways listed in > Table 2, using the corresponding representative gene sequences, an unequal distribution of these markers among phyla and genera is easily noticed. Only members of 8 out of 17 phyla where representatives have been sequenced show the presence of the catabolic gene markers described above. However, it should be also noted that among the phyla showing absence of aromatic catabolic pathway markers, often only a few representatives have been sequenced, such as one Verrucomicrobia, two Aquificae, Fusobacteria, or Lentispharea strains, three

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. Table 2 Key groups of catabolic enzymes discussed in the metabolic diversity section Enzyme group

Family

Pathway marker

Enzyme function

Abbreviation

Protocatechuate 3,4dioxygenase

Intradiol dioxygenase

++

Intradiol cleavage

Pca34

Catechol 1,2-dioxygenase

Intradiol dioxygenase

++

Intradiol cleavage

Cat12

Hydroxybenzoquinol 1,2dioxygenase

Intradiol dioxygenase

++

Intradiol cleavage

Hqu

Chlorocatechol 1,2-dioxygenase

Intradiol dioxygenase

+

Intradiol cleavage

Cca

Catechol 2,3-dioxygenase

Type I extradiol dioxygenase

++

Extradiol cleavage

Cat23

2,3-Dihydroxybiphenyl 1,2-dioxygenase

Type I extradiol dioxygenase

#

Extradiol cleavage

Dhb

Homoprotocatechuate 2,3-dioxygenase

Type I extradiol dioxygenase

++

Extradiol cleavage

HpcEXDOI

Protocatechuate 4,5dioxygenase

LigB-type dioxygenase

++

Extradiol cleavage

Pca45

Gallate 4,5-dioxygenase

LigB-type dioxygenase

++

Extradiol cleavage

Gal

Homoprotocatechuate 2,3-dioxygenase

LigB-type dioxygenase

++

Extradiol cleavage

HpcLigB

2,3-Dihydroxyphenylpropionate 1,2-dioxygenase

LigB-type dioxygenase

++

Extradiol cleavage

Dhp

2-Aminophenol 1,6-dioxygenase LigB-type dioxygenase

+

Extradiol cleavage

Amn

Gentisate 1,2-dioxygenase

Cupin superfamily dioxygenase

++

Extradiol cleavage

Gen

Homogentisate 1,2-dioxygenase Cupin superfamily dioxygenase

++

Extradiol cleavage

Hge

3-Hydroxyanthranilate 3,4-dioxygenase

Cupin superfamily dioxygenase

++

Extradiol cleavage

Han

Benzoquinol 1,2-dioxygenase

Type IV extradiol dioxygenase

+

Extradiol cleavage

Bqu

Benzoyl-CoA oxygenase

Soluble diiron oxygenase

++

Dearomatization

Box

Phenylacetyl-CoA oxygenase

Soluble diiron oxygenase

++

Dearomatization

Paa

2-Aminobenzoyl-CoA monooxygenase/reductase

Class A flavoprotein monooxygenase

++

Dearomatization

Abc

4-Hydroxybenzoate 3-hydroxylase

Class A flavoprotein monooxygenase

Forming protocatechuate

Phb3H

3-Hydroxybenzoate 4-hydroxylase

Class A flavoprotein monooxygenase

Forming protocatechuate

Mhb4H

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. Table 2 (Continued) Enzyme group

Family

Pathway marker

Enzyme function

Abbreviation

Salicylate 1-hydroxylase

Class A flavoprotein monooxygenase

Forming catechol

Ohb1H

3-Hydroxybenzoate 6-hydroxylase

Class A flavoprotein monooxygenase

Forming gentisate

Mhb6H

3-Hydroxyphenylpropionate 2-hydroxylase

Class A flavoprotein monooxygenase

Forming 2,3Mhp2H dihydroxyphenylpropionate

3-Hydroxyphenylacetate 6-hydroxylase

Class A flavoprotein monooxygenase

Forming homogentisate

Mha6H

Phenol/benzoquinol hydroxylase

Class A flavoprotein monooxygenase

Forming catechol/ hydroxybenzoquinol

Pbq2H

4-Hydroxyphenylacetate 3-hydroxylases

Class D flavoprotein monooxygenase

Forming homoprotocatechuate

Pha3H

Phenol 2-hydroxylase

Class D flavoprotein monooxygenase

Forming catechol

Ph2H

Chlorophenol 4-hydroxylase

Class D flavoprotein monooxygenase

Forming chlorobenzoquinol Ph4H

4-Hydroxyphenylacetate 3-hydroxylases

Class D* flavoprotein monooxygenase

Forming homoprotocatechuate

Pha3H

Resorcinol 4-hydroxylase

Class D* flavoprotein monooxygenase

Forming hydroxybenzoquinol

Res4H

Terephthalate 1,2-dioxygenase

Rieske nonheme iron oxygenase

Channeling to protocatechuate

TphDO

Phthalate 3,4-dioxygenase

Rieske nonheme iron oxygenase

Channeling to protocatechuate

Pht34DO

Anthranilate 1,2-dioxygenase (2 component)

Rieske nonheme iron oxygenase

Channeling to catechol

AntDO

Anthranilate 1,2-dioxygenase (3 component)

Rieske nonheme iron oxygenase

Channeling to catechol

AntDO

Benzoate 1,2-dioxygenase

Rieske nonheme iron oxygenase

Channeling to catechol

BenDO

Salicylate 5-hydroxylase

Rieske nonheme iron oxygenase

Channeling to gentisate

Sal5H

Phenylpropionate 2,3-dioxygenase

Rieske nonheme iron oxygenase

Channeling to 2,3PhpDO dihydroxyphenylpropionate

Biphenyl 2,3-dioxygenase type

Rieske nonheme iron oxygenase

Activation of hydrophobic aromatics

BphDO

Naphthalene inducible dioxygenase (NidA) type

Rieske nonheme iron oxygenase

Polycyclic aromatic degradation

NidDO

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. Table 2 (Continued) Enzyme group

Family

Pathway marker

Enzyme function

Abbreviation

Phthalate dioxygenase 4,5-dioxygenase

Rieske nonheme iron oxygenase

Channeling to protocatechuate

PhtDO

Isophthalate dioxygenase

Rieske nonheme iron oxygenase

Channeling to protocatechuate

IphDO

Toluene/benzene monooxygenase

Soluble diiron oxygenase

Forming phenol

Tmo

Phenol monooxygenase

Soluble diiron oxygenase

Forming catechol

Pmo

Only enzymes where a function could be assigned with high probability are included in the list Eleven groups of aromatic ring-cleavage activities (homoprotocatechuate 2,3-dioxygenases, even though belonging to different enzyme families were defined as one activity) and all groups of enzymes catalyzing dearomatization of aromatic CoA derivatives were defined as abundant, as they are observed in more than ten sequenced genomes and are marked as ++ Three groups of aromatic ring-cleavage activities were defined as less abundant, as they were observed in ten or less sequenced genomes and are marked as + 2,3-Dihydroxybiphenyl 1,2-dioxygenases (marked #) are not included in the list of aromatic catabolic pathway markers discussed in the proteobacterial section, as they are assumed to have their function in the metabolism of bi- and polycyclic aromatics rather than monocyclic aromatics Class D* flavoprotein monooxygenases refers to enzymes capable of using FMN, FAD, and riboflavin for hydroxylation

Planctomycetes, six Thermotogae, or nine Spirochaetes. Specifically in case the phylum contains aerobic species, only further genome analysis will reveal if such capabilities are in fact absent. Aromatic metabolic pathways were also absent from Chlamydiaea (11 genomes) where cultured representatives are obligate intracellular parasites of eukaryotic cells, the typically strict anaerobic Chlorobi (10 genomes), but also from Cyanobacteria (40 genomes), even though, for example, phenol degradation by the cyanobacterium Phormidium valderianum has been reported (Shashirekha et al., 1997). Most of the catabolic markers analyzed here are exclusively observed in Proteobacteria and Actinobacteria. This may be due to the fact that an immense amount of work has been invested specifically on elucidation of aromatic degradation in easy to culture members of these phyla. It thus cannot be excluded that novel groups of catabolic enzymes will be identified from other phyla. However, members of certain catabolic gene families can be observed in some representatives of other genera, such that the genome survey performed here is valid to get a reasonable overview of metabolic properties also from other phyla. For example, members of the cupin family, i.e., gentisate 1,2-dioxygenase, homogentisate 1,2-dioxygenase, and 3-hydroxyanthranilate 3,4-dioxygenase are all observed in other phyla, with homogentisate 1,2-dioxygenase being observed in Bacteroidetes, Chloroflexi, and Firmicutes (Bacilli). Bacilli and Bacteroidetes were also indicated not only to encode gentisate 1,2-dioxygenase and 3-hydroxyanthranilate 3,4-dioxygenase, but also a phenylacetate degradative pathway. In contrast to ring-cleavage pathways mediated by members of the cupin family, pathways mediated by other extradiol dioxygenases or intradiol dioxygenases are scarce outside of the Actinobacterial and Proteobacterial phyla. Intradiol cleavage dioxygenases are observed in Acidobacteria and the Thermus/Deinococcus phylum, among the LigB-type extradiol dioxygenases only

1381

1382

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Phylogenomics of Aerobic Bacterial Degradation of Aromatics

homoprotocatechuate 2,3-dioxygenases is observed in Bacilli and out of EXDO I proteins only homoprotocatechuate 2,3-dioxygenase is observed in Bacilli and Thermus/Deinococcus. Exceptional is the detection of distinct EXDO I proteins in Chloroflexi. Even though only two Acidobacteria and four Deinococcus/Thermus strains have been sequenced, the genomic survey indicates aromatic metabolic properties to be spread among those phyla. It can be suggested that S. usitatus Ellin6076 is capable to degrade 4-hydroxybenzoate via protocatechuate followed by intradiol cleavage and 4-hydroxyphenylpyruvate via the homogentisate pathway. Further capabilities of Acidobacteria thus remain to be discovered. All four members of the phylum Deinococcus/Thermus obviously share the capability to degrade 4-hydroxyphenylacetate via homoprotocatechuate and D. geothermalis DSM 11300 seems to harbor the capability to degrade 4-hydroxybenoate via protocatechuate and intradiol cleavage. Intradiol cleavage seems to be absent from Chloroflexi, Bacteroidetes, and Firmicutes. Interestingly, Chloroflexi can be proposed to be phenol degraders catabolizing it via catechol and meta-cleavage. Among Bacteroidetes, the homogentisate pathway and astonishingly the 3-hydroxyanthranilate pathway, in addition to the phenylacetate degradative pathway, seem to be spread among members of the orders Flavobacteriales and Sphingobacteriales. Out of the Firmicutes, only Bacillaceae (members of the genera Bacillus, Exiguobacterium, Geobacillus, and Oceanobacillus have been sequenced) seem to harbor aromatic metabolic properties. Unfortunately, no Paenibacillus genome sequence is available so far. Bacillus strains such as Bacillus sp. JF8 (Shimura et al., 1999), B. subtilis IS13 (Shimura et al., 1999), and others have been shown to be capable of degrading aromatics such as biphenyl, guaiacol, cinnamate, coumarate, or ferulate (Peng et al., 2003), and Paenibacilli such as P. naphthalenovorans, Paenibacillus sp. strain YK5, or Paenibacillus sp. KBC101 (Daane et al., 2002; Iida et al., 2006; Sakai et al., 2005) are shown to be capable of degrading naphthalene, dibenzofuran, or biphenyl. Thus, the metabolic diversity of Bacillaceae is clearly underrepresented by the currently sequenced 28 genomes, which indicate metabolic properties similar to those of Bacteroidetes, such as a spread of the homogentisate pathway in Bacillus and the presence of the 3-hydroxyanthranilate and the gentisate pathway in addition to the phenylacetate degradative pathway in members of different genera. In addition, 4-hydroxyphenylacetate degradation via homoprotocatechuate seems to be also a capability spread among Bacillaceae.

5.2

Actinobacteria

Aromatic metabolic routes can be observed in 12 out of 20 families from the phylum Actinobacteria and pathways analyzed here are absent in Actinomycetaceae, Cellulomonadaceae, Kineosporiaceae, Microbacteriaceae, Nocardiopsaceae, Propionibacteriaceae, Bifidobacteriaceae, and Coriobacteriaceae. Within the Corynebacterium genus, C. diphteriae and C. jeijekum, a nocosomial pathogen have no aromatic catabolic pathways. Interestingly, they have the smaller genomes of this group. A similar situation is observed within the Mycobacteria, as M. leprae, M. bovis, and M. tuberculosum also have no aromatic catabolic pathways and the smaller genomes of this group. In contrast, environmental Mycobacteria are characterized by an enormous metabolic potential, however, it should be noted that M. vanbaalenii Pyr1, M. gilvum PYR-GCK, as well as strains JLS, KMS, and MCS have been sequenced due to their capability to degrade various polycyclic aromatics reflected in the presence of up to four NidA-type Rieske non-heme iron oxygenases for initiating metabolism of PAHs and up to six BphC type I extradiol dioxygenases per genome.

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

However, not only Mycobacteria are endowed with a high metabolic potential. In contrast to members of all phyla described above, Actinobacteria not only often comprise a homogentisate pathway, which is observed in seven families, but also a protocatechuate intradiol cleavage pathway observed in eight families and more than one third of sequenced strains. Typically, actinobacterial strains endowed with a protocatechuate pathway also harbor a protocatechuate forming 4-hydroxybenzoate 3-hydroxylase such as both Micrococcaceae or Streptomycetaceae (see > Table 3) and often a 3-hydroxybenzoate 4-hydroxylase (such as both Micrococcaceae), indicating protocatechuate to be a central intermediate of various metabolic routes. Interestingly, Mycobacteria harboring a protocatechuate intradiol cleavage do not contain any of the aforementioned genes, but typically a phthalate dioxygenase. > Table 3 shows an overview of catabolic markers observed at least twice in genomes of actinobacterial families, from which at least two genomes have been sequenced. Two observations are evident from the table. First, Corynebacteriaceae, Nocardiaceae, and specifically Micrococcaceae are endowed with a broad metabolic potential. However, it should be noted that among the three Nocardiaceae, R. jostii RHA1 has a metabolic potential much broader than Nocardia farcinica IFM 10152 or Nocardioides sp. JS614. Unfortunately, no more sequences of the reported highly versatile Rhodococcus genus (van der Geize and Dijkhuizen, 2004) are available thus far. Also various reports on the metabolic versatility of Arthrobacter strains are known (Nordin et al., 2005). In contrast, Corynebacteria just recently have become the focus of more intense metabolic investigations (Huang et al., 2006). Second, the table shows a clear cooccurrence of ring-cleavage activity markers as well as of markers for peripheral activities, supporting that our annotation efforts are appropriate to deduce metabolic potential.

5.3

Proteobacteria

Three of the five classes of Proteobacteria (a, b, and g) concentrate the vast majority of the reported catabolic pathways towards aromatic compounds that can be traced in the current genome databases (> Table 4). Only a couple of aromatic catabolic pathways (Pca34, Hge, and Han) are found in some strains of the Myxococcales order of d proteobacteria and none in the e proteobacterial class. The a class of Proteobacteria has an uneven distribution of aromatic catabolic gene markers. None of the members of the three families of the order Rickettsiales have such catabolic properties. The small genome size of these members may be related to this trait. Aromatic ringcleavage pathways are also absent from all members of the Parvularculaceae, Bartonellaceae, and Erythrobacteraceae families and some members of the Aurantimonadaceae, Bradyrhizobiaceae, Methylobacteriaceae, Phyllobacteraceae, Rhodobacteraceae, Acetobacteracea, Rhodospirillaceae, and Sphingomonadaceae families. In contrast, four a- proteobacterial strains (Bradyrhizobium sp. BTAi1, S. wittichii RW1, Sagittula stellata E-37, and Silicibacter pomeroyii DSS-3) have 8–9 out of the 14 main pathways and another three strains (Bradyrhizobium japonicum USDA110, Bradyrhizobium sp. ORS278, and Jannaschia sp. CCS1) have seven main aromatic catabolic pathways suggesting Bradyrhizobium strains to be metabolically highly versatile. The most broadly distributed pathways in the a class of proteobacteria are Pca34 and Hge being observed in 30–40% of the sequenced genomes and in 11 and 9 families, respectively. Some catabolic pathways are only seldomly found in members of this proteobacterial class, and only N. aromaticivorans DSM 12444 has the Cat23 pathway and only X. autotrophicus Py2

1383

Pca34

Phb3H

++

Streptomycetaceae (2)

Pht34DO

++

+

+

+

TphDO +

+

Cat12 ++

+

+

++

BenDO +

++

+

+

+

++

++

+

++

++ +

+

HppDO ++

++

++

+

Mha6H +

Dhp +

+

+

Mhp2H +

HpcEXDOI +

(+)

+

Pha3H +

(+)

+

NidDO +

++

++

Dhb

++

+

++

++

+

++

+

++; More than 60% of the sequenced genomes of these proteobacterial families comprise a gene encoding the mentioned activity (number of sequenced representatives is given in parentheses); +, between 20 and 60%; (+), less than 20%. For abbreviations, see > Table 2. Only families where at least two members have been sequenced are included in the analysis

Bifidobacteriaceae (4)

Nocardioidaceae (3)

+

++

+

Gen

+

+

+

Mycobacteriaceae (17)

+

++

++

Mhb6H ++

++

++

Hge

Micromonosporaceae (2)

Micrococcaceae (2)

Microbacteriaceae (2)

Frankiaceae (3)

Corynebacteriaceae (5)

Cellulomonadaceae (2)

Actinobacterial Families

Mhb4H

. Table 3 Catabolic gene markers of Proteobacteria

Paa

39 Abc

1384 Phylogenomics of Aerobic Bacterial Degradation of Aromatics

(+)

+

Pht34DO

(+)

TphDO

(+)

Cat23 (+)

Ohb1H (+)

AntDO ++

(+)

PMO (+)

(+)

Ph2H (+)

(+)

+

(+)

(+)

(+)

+

(+)

+

++

++

+

++

(+)

(+)

(+)

+

(+)

+

++

(+)

+

(+)

(+)

Comamonadaceae (8)

++

+

(+)

++

Burkholderiaceae (43)

(+)

++

Alcaligenaceae (3)

β Proteobacterial Families

(+)

+

++ +

(+) (+)

++ (+)

(+)

+

+

++

+

+

+

(+)

++

++

++

++

++

++

+

+

++

++

++

+

++

(+)

+

++

+

+

+

++

Sphingomonadaceae (5)

(+)

(+)

+

(+)

++

++

HppDO

Erythrobacteraceae (3)

SAR11 (2)

Rickettsiaceae (16)

Anaplasmataceae (10)

++

(+)

+

BenDO +

(+)

Pbq2H

+

Rhodospirillaceae (3)

++

++

Cat12

+

TMO (+)

Hqu

Acetobacteraceae (3)

++

Xanthobacteraceae (2)

++

+

(+)

Res4H

+

++

Rhizobiaceae (5)

++

+

+

Bqu

++

++

Phyllobacteriaceae (3)

++

++

Gen +

Mhb6H

Rhodobacteraceae (24)

+

Methylobacteriaceae (3)

Pca45

+

Mhb4H

+

Ohb5H +

++

Brucellaceae (6)

++

Phb3H

+

Hge

Hyphomonadaceae (3)

(+)

Bradyrhizobiaceae (11)

Bartonellaceae (3)

++

+

Aurantimonadaceae (2)

Pca34

Caulobacteraceae (2)

α Proteobacterial Families

IphDO

. Table 4 Catabolic gene markers of Actinobacteria Mha6H +

(+)

(+)

(+)

Han (+)

(+)

(+)

(+)

(+)

Dhp (+)

+

Mhp2H (+)

Gal (+)

HpcLigB ++

+

(+)

++

(+)

+

HpcEXDOI +

+

+

+

Pha3H (+)

+

Paa +

++

++

(+)

+

+

+

+

++

Box ++

(+)

++

(+)

(+)

Abc ++

+

+

(+)

+

+

+

+

+

+

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39 1385

PhpDO

Mhb4H

+

AntDO (+)

(+)

Hqu +

Bqu (+)

(+)

(+)

+

+

+

+

Res4H

TphDO

Pht34DO

(+)

+ +

(+)

(+)

+

(+) ++

++

++

++

++

++

+

+

+

(+)

+

+

+

++

++

(+)

(+)

+

+

(+)

+

(+)

+

(+)

+

(+)

+

+

+

(+)

++

(+)

++

Box

Mha6H

++

++; More than 60% of the sequenced genomes of these Actinobacterial families comprise a gene encoding the mentioned activity (number of sequenced representatives is given in parentheses); +, between 20 and 60%; (+), less than 20%. For abbreviations, see > Table 2. Only families where at least two members have been sequenced are included in the analysis

++

+

++

++

+

Xanthomonadaceae (11)

+

++

+

Vibrionaceae (30)

Thiotrichaceae (2)

+

++

Pseudomonadaceae (19)

Francisellaceae (7)

+

Moraxellaceae (5)

Pasteurellaceae (21)

++ +

++

+

Oceanospirillaceae (3)

+

Legionellaceae (4)

Coxiellaceae (5)

Enterobacteriaceae (61)

+

Han (+)

Mhp2H

(+)

+

++

+

Dhp ++

PhpDO

+

(+)

HppDO +

Gal

Ectothiorhodospiraceae (3)

(+)

Ph2H (+)

+

HpcLigB

+

(+)

(+)

Pbq2H +

HpcEXDOI

++

(+)

Ohb1H (+)

(+)

PMO +

Pha3H

Shewanellaceae (18)

(+)

(+)

TMO +

Paa

Psychromonadaceae (2)

(+)

Cat12

+

BenDO +

Gen

++

+

IphDO

(+)

Cat23 +

Mhb6H

Pseudoalteromonadaceae (3)

+

Phb3H

+

Ohb5H ++

(+)

Pca45

+

(+)

Hge

Idiomarinaceae (2)

Aeromonadaceae (7)

γ Proteobacterial Families

Rhodocyclaceae (3)

Nitrosomonadaceae (3)

Neisseriaceae (5)

39

Oxalobacteraceae (2)

α Proteobacterial Families

Pca34

. Table 4 (Continued) Abc

1386 Phylogenomics of Aerobic Bacterial Degradation of Aromatics

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

39

has the Dhp pathway. The Cca, Amn, and Bqu pathways are not found in any a proteobacterial genome. Regarding peripheral pathways, a proteobacterial strains endowed with a protocatechuate pathway also harbor a Phb3H and with lower frecuency a Mhb4H. Isomers of phthalate seems not to be typical substrates for a proteobacteria, since with the exception of IphDO in B. japonicum USDA110, phthalate, isophthalate, or terephthalate dioxygenases are not found. BenDO are usually observed in strains endowed with a Cat12 pathway and strains endowed with Hge usually also harbor HppDO encoding genes. The b class of proteobacteria harbors all major central aromatic catabolic pathways listed in > Table 2. The distribution of these catabolic pathways among b-proteobacterial strains has some points to be noted. Except for the presence of the Hge catabolic pathway in C. violaceum, the families Oxalobacteraceae, Neisseriaceae, and Nitrosomonadaceae are devoid of the investigated aromatic catabolic properties. Specifically, members of the Burkholderiaceae and Comamonadaceae show a high metabolic potential and usually harbor a broad set of aromatic pathways and members of the Burkholderia, Cupriavidus, Ralstonia, Delftia, and Polaromonas genera comprise up to 11 out of the 14 major central aromatic pathways (Pe´rezPantoja et al., 2008). Polynucleobacter sp. QLW-P1DMWA-1 is the only member of the Burkholderiaceae family that has no such catabolic pathway (the smallest genome among them); and Limnobacter sp. MED105 (the second smallest genome) has only Cat23. Members of the Alcaligenaceae and Rhodocyclaceae are obviously relatively limited in their aromatic catabolic potential. It should be noted that Rhodocyclaceae comprise genera such as Azoarcus, Thauera, or ‘‘Aromatoleum,’’ nitrate-reducing bacteria that contribute significantly to the biodegradation of aromatic compounds in anoxic waters and soils and that are endowed with several pathways for anaerobic catabolism of aromatics. It has, however, also been shown that aerobic aromatic pathway are functional in these bacteria (Rabus, 2005). The most abundant pathways in the b class are Paa, Hge, Cat12, and Pca34, which are found in 60% or more of the sequenced genomes available. In contrast, Gal, Dhb, and Han are only observed in Table 2, respectively, being among the catabolically most versatile bacteria reported so far. The inspection of the genome database allows the finding of other, sometimes unexpected, bacteria with broad aromatic metabolic potential. All three Cupriavius strains contain 9–11 of the main pathways and among Burkholderia strains, B. phymatum STM815, and Burkholderia sp. 383 deserve special attention, as they both contain ten of the main and one of the rare pathways. Also Bradyrhizobia, for which three genomes are available can be regarded as exceptionally versatile comprising seven to eight major pathways, as is also the case for some marine Rhodobacteraceae (Silicibacter pomeroyi DSS-3 [Moran et al., 2004], Sagittula stellata E-37 [Gonzalez et al., 1997], Jannaschia sp. CCS1), Azoarcus sp. BH72 and various Comamonadaceae or Burkholderiaceae. These bacteria are, therefore, choices to perform metabolic reconstruction studies as indicated above, in order to demonstrate their catabolic potential. Interestingly, Pseudomonas strains only comprise up to six main metabolic pathways, with P. putida W619 isolated from the Black Cottonwood tree having the broadest metabolic potential. It can be assumed that these highly versatile catabolic bacteria live in environments where a variety of aromatic carbon sources are present. One kind of such habitats is the rhizosphere of plants, since it is expected that their exudates contain a myriad of organic carbon sources, most of them in tiny amounts. Interestingly, several of the bacteria listed above have been isolated or proposed to thrive in rhizospheric habitats. Even more, some of them have been described to produce beneficial effects on plants (Bradyrhizobium, Burkholderia) suggesting a mutually positive interaction between plants and versatile aromatic degraders.

6

Research Needs

Hundreds of bacterial genomes have been completely sequenced, several of which are important paradigms for pollutant transformation pathways. Such complete information of bacterial cells will allow in concert with transcriptomic and proteomic studies the analysis of the detailed behavior and physiology of these organisms, the development of bioinformatic models, and also a predictive modeling. However, various other aspects have to be considered

1391

1392

39

Phylogenomics of Aerobic Bacterial Degradation of Aromatics

to reach such goals. First, misannotations in bacterial genome projects are too frequent. However, the identification in databases of proteins of which a function has been proven, to allow a comparison with the protein of interest becomes more and more complicated with the overwhelming data arising from the sequencing projects. Cured databases with valid information are necessary, such as the ribosomal database project (RDP) at rdp.cme.msu.edu or the TCDB transport classification database at www.tcdb.org, which have been developed to facilitate analysis of 16S rDNA or membrane transport proteins. Second, the amount of genes coding for proteins with unknown function is immense, and even broader than annotations may suggest. Metabolic reconstruction work may help to elucidate metabolic routes for which the genetic basis has not yet been explored (Nogales et al., 2008; Pe´rez-Pantoja et al., 2008). In this context, it is also important to note that an immense amount of valuable information is available on the biochemistry of metabolic pathways, and even though mainly dating back 30 or more years ago is a source that should be recommended for reading. Even though the list of bacterial genomes sequenced or in the process of sequencing is enormous, the current database is still highly biased for easy to culture microorganisms, as expected, and even some environmentally important groups such as Rhodococcus sp. are represented just by one genome. Efforts should be directed towards a better understanding of the diversity inside such genera. However, it should be also noted that significant efforts are needed to really harvest the information available from already sequenced genomes.

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Asturias JA, Timmis KN (1993) Three different 2,3dihydroxybiphenyl-1,2-dioxygenase genes in the gram-positive polychlorobiphenyl-degrading bacterium Rhodococcus globerulus P6. J Bacteriol 175: 4631–4640. Ballou DP, Entsch B, Cole LJ (2005) Dynamics involved in catalysis by single-component and two-component flavin-dependent aromatic hydroxylases. Biochem Biophys Res Commun 338: 590–598. Beil S, Mason JR, Timmis KN, Pieper DH (1998) Identification of chlorobenzene dioxygenase sequence elements involved in dechlorination of 1,2,4,5tetrachlorobenzene. J Bacteriol 180: 5520–5528. Beltrametti F, Marconi AM, Bestetti G, Colombo C, Galli E, Ruzzi M, Zennaro E (1997) Sequencing and functional analysis of styrene catabolism genes from Pseudomonas fluorescens ST. Appl Environ Microbiol 63: 2232–2239. Bosch R, GarciaValdes E, Moore ERB (1999a) Genetic characterization and evolutionary implications of a chromosomally encoded naphthalene-degradation upper pathway from Pseudomonas stutzeri AN10. Gene 236: 149–157. Bosch R, Moore ERB, GarciaValdes E, Pieper DH (1999b) Nah W, a novel, inducible salicylate hydroxylase involved in mineralization of naphthalene by Pseudomonas stutzeri AN10. J Bacteriol 181: 2315–2322.

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40 Transcriptional Networks that Regulate Hydrocarbon Biodegradation G. Carbajosa . I. Cases Spanish National Cancer Research Centre, Madrid, Spain [email protected] [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1400 2 General Patterns in Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1400 3 Specific Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1400 4 Complex Regulatory Networks and Cross-Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1402 5 Global Regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1403 6 Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1405 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1406

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_96, # Springer-Verlag Berlin Heidelberg, 2010

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Transcriptional Networks that Regulate Hydrocarbon Biodegradation

Abstract: Transcription Regulation is one of the key steps in hydrocarbon biodegradation. The expression of the genes required for hydrocarbon degrading pathways can be tightly regulated in response to substrate availability and physiological status of the cells. Also different pathways have to be properly coordinated to ensure maximal efficiency in the use of available nutrients. Different mechanisms have been developed by bacteria to achieve this regulation, which vary in complexity and in the involved elements. In this chapter, we summarize these mechanisms, giving recent examples, and describe some of the interesting applications that the engineering of regulatory circuits can bring to the fields of bioremediation.

1

Introduction

Since the industrial revolution, human activities have resulted in the release and introduction of several kinds of aromatic compounds, displaying more novel structures than that already present in natural environments. In response to those environmental insults, microorganisms and microbial communities have developed the ability to process such recalcitrant compounds that do not form part of their central metabolism by transformations, that interestingly, lead to the final introduction into it. These new acquired capabilities are of great interest and there have been several attempts to use them in order to deal with environmental catastrophes and pollution in general (Cases and de Lorenzo, 2005b). But the presence of biodegradation abilities does not guarantee the use of them. Microorganisms have a tight regulation that allows them to express catabolic genes under a changing environment (Cases and de Lorenzo, 2005a) integrating the signals with the cell physiology (Cases and de Lorenzo, 2001). In a way, transcriptional regulation acts as a limiting factor that restricts the production of these secondary metabolism enzymes, which degrade hard palatable compounds, allowing their expression only when no other carbon source is available or in response to a particular stress (Cases and de Lorenzo, 2005a). Therefore, the study of the regulatory systems implicated in the expression of xenobiotic catabolic pathways is of main interest if we want to unravel the mechanism behind the biodegradation processes and understand how microorganisms behave in their natural environments.

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General Patterns in Regulation

Transcriptional regulation of catabolic pathways acts at two levels: the specific level, where a regulator induces transcription initiation of the catabolic genes by interacting with the aromatic substrate of the pathway, or a related compound, and the global level, where the expression of catabolic genes is integrated with the host physiology.

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Specific Regulation

Specific regulation of catabolic pathways generally involved the participation of a transcription factor that upon interaction with a small molecule, induce the expression of the genes encoding the required enzymes. While this small molecule is often the main substrate of the pathways, in some systems, an intermediate compound of the pathway is selected as the best inducer. This is the case of BenM in Acinetobacter baylyi ADP1 that is induced by cis,

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cis-muconate (Ezezika et al., 2007a). This is being suggested to be the best compromise position in terms of robustness and responsiveness for these inducible regulatory systems (Wall et al., 2004). Surprisingly, the resolution of the crystallized structure of this regulator revealed the presence of two effector binding sites (Ezezika et al., 2007a) responsible for a previously described synergistic effect: the expression of its regulated promoter is significantly higher in response to cis,cis-muconate and benzoate than the combined levels due to each alone (Bundy et al., 2002). This fact suggests a new direction in the study of the aromatic regulatory systems, as this dual binding could not be an isolated case. Interestingly, biodegradation-related transcription factors often interact with a broad range of chemical compounds, and while most of theses compounds are structurally related to the natural substrates of the regulated pathway, some are not. This ability of regulators under control of aromatic biodegrading pathways to respond to several compounds is widely spread. VanR, a GntR-type regulator found in Caulobacter crescentus, was described to respond to several compounds but vanillate was found to be the best inducer (Thanbichler et al., 2007). Catechol, 2-methylhydroquinone and chromanon induce YodB, a Bacillus subtilis MarR repressor. In Pseudomonoas fluorescens ST, a range of compounds induces the TCS StyRP but only Toluene generates a high response. This capability of a regulator to respond to a wide range of compounds has been previously described and has been suggested to allow systems to evolve and acquire the capability to respond to novel environmental signals (regulatory noise): ‘‘Transcriptional control systems develop responsiveness to new signals due to the leakiness and lack of specificity of preexisting promoters and regulators. When needed, these may become more specific through suppression of undesirable signals and further fine-tuning of the recruited proteins to interact with distinct chemicals’’ (de Lorenzo and Perez-Martin, 1996). However, it was recently found that transcription factors can also be coupled to the regulated pathways be other means. A model for the degradation of tetralin has been proposed where a direct communication between its regulatory system and ferredoxin, coded by thnA3, in its reduced form, prevents induction of the system by a molecule that is not a real substrate. This communication allows a better fit of the substrate and inducer profiles, thus minimizing gratuitous induction, without a requirement for optimal coevolution to match the specificity of catabolic enzymes and their regulatory systems. Modulation of the regulatory system in this way not only provides a more appropriate response to potential inducers recognized by the regulatory system but also may properly adjust the levels of gene expression to the substrate availability (Martinez-Perez et al., 2007). This model opens a new approach to the study of aromatic catabolic pathways regulation that is yet unexplored. Aromatic catabolic pathways are not controlled by a single type of regulators but by several different families of regulators that are not specific to biodegradation processes. The different regulatory families involved in aromatic compound biodegradation include LysR, IclR, AraC/ XylS, GntR, TetR, MarR, FNR, two-component regulatory systems (TCSs), XylR/NtrC, and the recently added LuxR, PadR, and SinR. Interestingly, there is no relation between the transcription factor family and the compound they interact with. For example, a carbazole degradation id regulated by CarRI, an IclR-like regulator in Nocardioides aromaticivorans (Inoue et al., 2006), by an AraC-type regulator, AntR, in Pseudomonas resinovorans CA10 (Miyakoshi et al., 2006; Urata et al., 2004), and by a GntR regulator, CarR in Janthinobacterium sp. J3 (Miyakoshi et al., 2006). For the catechol degradation we can mention two recent examples: CatR in Rhodococcus erythropolis CCM2595, an IclR-type transcriptional regulator (Vesely et al., 2007), and MhqR in Bacillus subtilis, a MarR-type regulator (Towe et al., 2007). A review comparing naphtalene-degrading clusters also shows up different types of regulators,

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from LysR, NtrC, and GntR families, for systems implicated in the degradation of such compound (Kulakov et al., 2005). To explain these broad spectrum of types of regulators, the hypothesis that catabolic and regulatory genes have evolved relatively independently has been proposed (Cases and de Lorenzo, 2001). If this is true, that would mean that the capability to respond to specific substrates, or related compounds, of the same catabolic pathway has been developed independently several times through evolution.

4

Complex Regulatory Networks and Cross-Regulation

While as described before, regulatory circuits of catabolic pathways tend to be simple, the distribution of the structural genes in two, or more operons, involving the presence of regulatory cascades is not uncommon. For instance, the TOL plasmid in Pseudomonas putida mt2 is the case best studied. An ‘‘upper’’ pathway is clustered in an operon under the control of XylR, an NtrC-type regulator, and the ‘‘lower’’ pathway hosted by another operon is regulated by XylS, an AraC regulator. XylS gene is regulated by XylR fine-tuning the system (Marques et al., 1998). Also, it is not unusual that organisms hosting several biodegradation pathways display interconnections between them. This interplay could be controlled through genetic organization, which is the case of protocatechuate pathway regulator PcaU in Acinetobacter. It has been discovered that it regulates quinate degradation too by transcribing the qui gene cluster, immediately downstream of the protocatechuate degrading genes, through a 14-kb transcript (Dal et al., 2005). When an organism has the capability of producing several degrading pathways, the presence of the different inducers that stimulate the expression of some of them can be resolved in different ways. In Acinetobacter baylyi ADP1, BenM, the regulator that activates benzoate degradation can be synergistically activated by the pathway substrate and cis,cis-muconate, a pathway intermediate metabolite. CatM, a BenM paralog, is activated by cis,cis-muconate too, but not by benzoate, and induces the expression of the catecholdegrading cluster. They both show cross-regulation. (Ezezika et al., 2006, 2007a, b). Furthermore, the expression of the regulator of protocatechuate degradation, PcaU, is repressed by either CatM or BenM in the presence of the pathway intermediate cis,cis-muconate, thus selecting the catechol-degrading branch and repressing the protocatechuate-degrading one (Brzostowicz et al., 2003). Some other studies describing cross-regulation of operons by different regulators also suggest that the regulation of catabolic promoters can be far more complex than the traditional couple operon-specific regulator approach. BphR2 is a new described regulator in Pseudomonas pseudoalcaligens KF707, which binding sequences exhibit similarity with those of NahR, which cross-regulates the biphenyl and salicylate catabolic genes in interplay with BphR. The presence of the substrates, or intermediate compounds, of the two catabolic clusters alters the expression of the regulators that bind to the operator regions, finetuning the system response by means of a regulatory cascade (Fujihara et al., 2006; Furukawa, 2006; Furukawa and Fujihara, 2008; Furukawa et al., 2004; Takeda et al., 2004). Examples of catabolic pathways coordination are becoming more abundant, although in many cases the molecular details remain unknown. For instance, Rhodococcus sp. DK17 requires that benzoate in the medium is depleted before the utilization of phthalate begins while the transcription of the genes encoding benzoate and phthalate dioxygenases parallel the substrate utilization in a catabolite repression-like response (Choi et al., 2007), while a new regulatory system found in Pseudomonas putida U, MhaSR, seems to be crucial for the hierarchical utilization of different

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aromatic compounds as carbon sources, as well as the flux of intermediates through the different pathways involved in the degradation of such compounds (Arias-Barrau et al., 2005).

5

Global Regulation

Since the discovery that the catabolic promoters expression is tightly connected with the host physiology (Cases and de Lorenzo, 2001, 2005a), the scientific community has made an effort to understand and explain the mechanisms performing such integration. For example, a tighter integration than what was previously known between the TOL-plasmid encoded functions and the host physiology has been described: posttranscriptional checks appear to mitigate the burden caused by nonproductive induction of the TOL operons, the fate of different segments of the polycistronic mRNAs from the upper and lower operons varies depending on the metabolism of its inducers, and finally, the heat shock response provoked by m-xylene interferes with the optimal expression of the operons (Velazquez et al., 2005). This stress response has been investigated further suggesting that adaptation to external insults precedes any significant expression of catabolic genes (Velazquez et al., 2006). The presence of some specific inducers can also subvert the lag starvation period in Nocardioides sp. JS614 (Mattes et al., 2007). Another example reassuring this suggestion is the studied response to toluene in Pseudomonas putida KT2440 where it is sensed as a stressor rather than as a nutrient (Dominguez-Cuevas et al., 2006). Integration with physiological cues can be performed by a number of different mechanisms (> Fig 1). Participation of DNA-binding transcription factor is one of them. One example could be AadR. HbaR, a FNR-type regulator, which in Rhdopseudomonas palustris, controls the expression of hbaA encoding 4-HBA-CoA ligase, the first enzyme for 4-HBA degradation. AadR, another FNR regulator, activates expression of hbaR in response to anaerobiosis and HbaR, in turn, activates expression of 4-hydroxybenzoate (4-HBA) degradation in response to 4-HBA as an effector molecule (Egland and Harwood, 2000). Other less characterized examples are the effect of TurA and PprA on s the Pu promoter of the TOL plasmid. The growth of the bacterial host at suboptimal temperatures resulted in a TurA-dependent increase of Pu repression. This is a novel physiological modulation of a promoter that is different to all the strictly physiological previously described (Rescalli et al., 2004). PprA is a LytTR-type response regulator that represses the expression from the TOL plasmid Pu promoter. It performs its repression by occupying the target-binding sites of its cognate activator XylR, in a suggested mutual exclusion mechanism. It has been argued by the authors of the study that the binding of PprA could help to anchor the TOL regulatory subnetwork to the wider context of the host transcriptome, thereby allowing the entry of physiological signals that modulate the outcome of promoter activity (Vitale et al., 2008). Other major player in physiological integration is the global regulatory protein Crc. Crc is involved in the repression of several catabolic pathways for sugars, hydrocarbons, and nitrogenated and aromatic compounds in Pseudomonas putida and Pseudomonas aeruginosa when other preferred carbon sources are present in the culture medium (catabolite repression), therefore modulating carbon metabolism (Ruiz-Manzano et al., 2005). Several experiments performed in isogenic Pseudomonas putida strains showed up that Crc is involved in the catabolic repression of the homogentisate pathway, benzoate and catechol degradation, and protocatechuate pathway too (Morales et al., 2004). In controls the benzoate degradation

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. Figure 1 Many features besides the binding of the specific regulator can influence the expression of a specific catabolic operon, in order to integrate the expression of the operon with the host physiology. For example, the binding of a regulatory complex to a third binding site can inhibit the expression of the catBCA operon when a big amount of regulator is being produced. We called this here a binding effect. Some promoters may need the presence of an integration host factor (IHF) to be able to recruit the RNA polymerase (RNAP) and start transcription. IHF can prevent promiscuous activation by other regulators besides providing physical contacts to the RNAP by bending DNA. The transcription initiation factor (s factor) enables specific binding of RNA polymerase to gene promoters. Different sigma factors are activated in response to different environmental conditions triggering the expression of its regulated ‘‘sigmulons.’’ ppGpp is an alarmone that is involved in the stringent response in bacteria, causing the inhibition of RNA synthesis. For instance, ppGpp directly stimulates transcription from the Po and Pu promoters, specifically regulated by XylR and DmpR, implicated in the degradation of toluene and phenol respectively. Crc is a global regulatory protein implicated in the regulation of several catabolic pathways. It has been suggested to act on translation by preventing the access of the ribosomes to the translational start of the mRNA.

pathway, limiting the expression of its specific regulator, BenR, and preventing full expression of the catabolic genes in this way. Crc directly inhibits the expression of the peripheral genes that transform benzoate into catechol (the ben genes), but its effect on genes corresponding to further steps of the pathway (the cat and pca genes of the central catechol and beta-ketoadipate pathways) is indirect, since these genes are not induced because the degradation intermediates, which act as inducers, are not produced. Crc inhibits the the translation of target genes by binding to 50 end of benR mRNA (Moreno et al., 2007). In a similar manner, Crc controls the

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expression of AlkS reducing the activation of the promoter expressing alkane-degrading enzymes (Moreno et al., 2007). Its level and activity vary according to growth conditions. Expression of crc is high during exponential growth and decreases when the growth rate declines before cells enter into stationary phase. In stationary-phase cells, transcript levels are three- to fourfold lower than those of cells in mid-exponential phase. When cells are grown in minimal salts medium with citrate as a carbon source, a substrate that does not induce catabolite repression on the alkane degradation pathway, expression of crc is two- to threefold lower than in LBgrown cells and also declines in stationary phase (Ruiz-Manzano et al., 2005). Other mechanisms for signal integration are more subtle. For instance, some bacterial promoters need the assistance of DNA-bending proteins, such as IHF, in order to assemble the machinery that would start transcription. The Integration Host Factor (IHF) is an asymmetric histone-like protein that binds and bends the DNA at specific sequences. IHF functions as an accessory factor in a wide variety of processes including replication, site-specific recombination, and transcription (Goosen and van de Putte, 1995). A large number of s54-dependent promoters require this site-specific DNA-bending protein, including Po and Pu promoters involved in degradation of xylene and toluene, and phenol respectively (de Lorenzo et al., 1991; Sze et al., 2001). Its function goes beyond providing close physical contact: it prevents promiscuous cross-activation by other s54-depent regulators (Perez-Martin and De Lorenzo, 1995) and helps to recruit the core polymerase-sigma54 complex with a distally located ‘‘UPlike’’ DNA element that is otherwise out of reach (Bertoni et al., 1998). A model has been proposed for styrene degradation in Pseudomonas where IHF plays a crucial role: the TCS StyRS changes the StyR-P (phosphorilated) levels, depending on StyS-P levels, vary according to the relative ratio of StyS-P and IHF, which in turn would determine variations of the promoter architecture and modulation of its activity (Leoni et al., 2007). Physiological control can finally be also mediated by small messenger molecules such us ppGpp. ppGpp is probably the most far-reaching bacterial global signaling molecule both through its direct effects mediated by signaling at the interface of the b and b0 subunits of RNA polymerase, and indirectly, through regulatory cascades as a result of its stimulatory effects on promoters that control other global regulators (Shingler, 2003). For example, ppGpp directly stimulates transcription from the s54-dependent Po and Pu promoters (Carmona et al., 2000). ppGpp and its cofactor DksA also influence the expression from s54-depent promoters by modulating the competitive association with sigma factors and reducing innate stability of RNA-polymerase complexes (Bernardo et al., 2006; Szalewska-Palasz et al., 2007). In this way, the transcription of the bacterial promoters is determined by the physiology of the cell, here represented by the availability of the different sigma factors, and the ppGpp effect on the RNApolymerase

6

Applications

Transcriptional regulation of the degradation of aromatic compounds has proved to be interesting enough to interact with other fields of study, attracting the use of innovative techniques and opening new research possibilities. Here is a brief comment on some of them. A new-born field, synthetic biology aims to design and construct biologic circuits in the way engineers construct electronic circuits (Tucker and Zilinskas, 2006). By combining ‘‘biological parts,’’ like promoters, regulators, and operons, the objective is to obtain systems optimized for a given purpose. In an example of the use of these techniques, the XylR regulator is evolved,

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by gene shuffling plus mutagenic PCR, to be optimally responsive to 1,2,4-trichlorobenzene, the major product of gamma-hexachlorocyclohexane (g-HCH) dechlorination. Then a combination of its recognized promoter Pu with lacZ and linA allows the detection of levels of b-galactosidase that are dependent on the g-HCH concentration. Therefore, by using the ability of a regulatory protein to recognize the product of a given reaction, this activity can be detected in a host using a ‘‘reporter gene’’ like lacZ. (Mohn et al., 2006). Further efforts in this direction have crystallized with the production of mimetic regulatory factors by aiming singlechain antibodies to XylR (Jurado et al., 2006), the detection of salicylate in ADP1 constructing a gene fussion between the induced genes and GFP-reporter gene (Huang et al., 2005; Huang et al., 2006), or tracing explosives in soil using the ability of XylR to detect 2,4 dinitrotoluene (Garmendia et al., 2008). The cross-talk between chemical sensed by transcriptional factors has opened the possibility of developing regulators that can be used as biosensors for environmentally relevant compounds (Galvao and de Lorenzo, 2006). A successful example on this area of research is the development of a dual bioreporter based on cross-activation by XylR and HbpR. HbpR regulated promoter was changed by site-directed mutagenesis in to XylR binding sites. Some mutants were efficiently activated by both HbpR and XylR, showing that promoters can be created that are permissive for both regulators. On the basis of these results, a dual-responsive bioreporter strain of P. azelaica was created, containing both XylR and HbpR, and activating luciferase expression from the same single promoter independently with m-xylene and 2-hydroxybiphenyl (Tropel et al., 2004).

7

Research Needs

In the recent years, the understanding of regulatory systems behind aromatic compounds degradation has been broadened. Bioinformatics, in combination with high-throughput techniques such as genome sequencing and proteomics, is playing an important role in how we acquire new information about biodegradation regulation. For instance, genomic and proteomic approaches were used to investigate phthalate and benzoate catabolism in Rhodococcus sp. Strain RHA1, a polychlorinated biphenyl-degrading actinomycete. Posterior analysis showed up the presence of regulatory and structural genes. Among them, a new regulator controlling a benzoate-degrading cluster, namely BenR, and Two IclR-type regulators, PcaR and PdaR, implicated in phtalate degradation were described. Interestingly, high redundancy of catabolic pathways and enzymes appears to be unique to RHA1 and may increase its potential to adapt to new carbon sources (Patrauchan et al., 2005). In another example, a combination of bioinformatic tools and a comparative genomic approach was used to identify and characterize a set of conserved DNA-binding transcriptional regulators in four corynebacterial genomes. Among this regulators, severeral were related to the regulation of the metabolism of aromatic compounds, including gentisate, phenol. protocatechuate, p-cresol, 4-hydroxybenzoate, benzoate, benzylalcohol, resorcinol 2,4-dihydroxybenzoate, vanilline, and vanillate (Brune et al., 2005). Also, the sequencing of the complete genome of Dehalococcoides strain CBDB1, which catabolize many of the most toxic and persistent chlorinated aromatics and aliphatics by reductive dechlorination and used for in situ bioremediation of contaminated sites, has revealed the presence of 32 reductive-dehalogenase-homologous (rdh) genes, possibly conferring on the bacteria an immense dehalogenating potential. Most rdh genes were

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associated with genes encoding transcription regulators such as two-component regulatory systems or transcription regulators of the MarR-type (Kube et al., 2005). This amount of information on regulation and metabolism of biodegradation pathways will allow a systems biology approach to its study. Previous efforts in that direction include the study of the microbial biodegradation network from a systems biology perspective as an efficient and integrated supra-metabolism, with properties similar to those that define metabolic networks in single organisms (Pazos et al., 2003). Minnesota University has built a database that contains information on microbial biocatalytic reactions and biodegradation pathways for primarily xenobiotic, chemical compounds (Ellis et al., 2006). We have made our own contribution creating a database with all the regulatory and metabolic information available at the moment of its construction that contains detailed molecular information like binding sites, regulatory complexes, and promoters, named Bionemo (in press). This kind of tools would be useful in the near future to study the evolution of the systems, reveal the singular features that make these regulatory systems different from others and understand the interaction between different strains as occurs in natural environments. The traditional approach studying a gene cluster regulated by a specific regulatory protein has been overtaken by a more subtle research that takes into account the commonly found phenomenon of cross-regulation and complex networks. A focus has been set in unraveling the integration between host physiology and specific regulation of particular pathways and today we know a lot more about this subject. However, we are still far from being able to model these processes and actually predict their behavior in the environment. The possibilities of a systematic approach by bioinformatics tools to the study of the field are still to be developed. Bioinformatics and Systems biology will need to work together to make reliable predictions on biodegradation regulation. Applications in the field of Synthetic Biology and their main objective of construction biological circuits will be feasible only once we understand the logic behind the regulatory components.

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Transcriptional Networks that Regulate Hydrocarbon Biodegradation Marques S, Gallegos MT, Manzanera M, Holtel A, Timmis KN, Ramos JL (1998) Activation and repression of transcription at the double tandem divergent promoters for the xylR and xylS genes of the TOL plasmid of Pseudomonas putida. J Bacteriol 180: 2889–2894. Martinez-Perez O, Lopez-Sanchez A, Reyes-Ramirez F, Floriano B, Santero E (2007) Integrated response to inducers by communication between a catabolic pathway and its regulatory system. J Bacteriol 189: 3768–3775. Mattes TE, Coleman NV, Chuang AS, Rogers AJ, Spain JC, Gossett JM (2007) Mechanism controlling the extended lag period associated with vinyl chloride starvation in Nocardioides sp. strain JS614. Arch Microbiol 187: 217–226. Miyakoshi M, Urata M, Habe H, Omori T, Yamane H, Nojiri H (2006) Differentiation of carbazole catabolic operons by replacement of the regulated promoter via transposition of an insertion sequence. J Biol Chem 281: 8450–8457. Mohn WW, Garmendia J, Galvao TC, de Lorenzo V (2006) Surveying biotransformations with a la carte genetic traps: translating dehydrochlorination of lindane (gamma-hexachlorocyclohexane) into lacZ-based phenotypes. Environ Microbiol 8: 546–555. Morales G, Linares JF, Beloso A, Albar JP, Martinez JL, Rojo F (2004) The Pseudomonas putida Crc global regulator controls the expression of genes from several chromosomal catabolic pathways for aromatic compounds. J Bacteriol 186: 1337–1344. Moreno R, Ruiz-Manzano A, Yuste L, Rojo F (2007) The Pseudomonas putida Crc global regulator is an RNA binding protein that inhibits translation of the AlkS transcriptional regulator. Mol Microbiol 64: 665–675. Patrauchan MA, Florizone C, Dosanjh M, Mohn WW, Davies J, Eltis LD (2005) Catabolism of benzoate and phthalate in Rhodococcus sp. strain RHA1: redundancies and convergence. J Bacteriol 187: 4050–4063. Pazos F, Valencia A, De Lorenzo V (2003) The organization of the microbial biodegradation network from a systems-biology perspective. EMBO Rep 4: 994–999. Perez-Martin J, De Lorenzo V (1995) Integration host factor suppresses promiscuous activation of the sigma 54-dependent promoter Pu of Pseudomonas putida. Proc Natl Acad Sci USA 92: 7277–7281. Rescalli E, Saini S, Bartocci C, Rychlewski L, De Lorenzo V, Bertoni G (2004) Novel physiological modulation of the Pu promoter of TOL plasmid: negative regulatory role of the TurA protein of Pseudomonas putida in the response to suboptimal growth temperatures. J Biol Chem 279: 7777–7784.

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Ruiz-Manzano A, Yuste L, Rojo F (2005) Levels and activity of the Pseudomonas putida global regulatory protein Crc vary according to growth conditions. J Bacteriol 187: 3678–3686. Shingler V (2003) Integrated regulation in response to aromatic compounds: from signal sensing to attractive behaviour. Environ Microbiol 5: 1226–1241. Szalewska-Palasz A, Johansson LU, Bernardo LM, Skarfstad E, Stec E, Brannstrom K, Shingler V (2007) Properties of RNA polymerase bypass mutants: implications for the role of ppGpp and its co-factor DksA in controlling transcription dependent on sigma54. J Biol Chem 282: 18046–18056. Sze CC, Laurie AD, Shingler V (2001) In vivo and in vitro effects of integration host factor at the DmpRregulated sigma(54)-dependent Po promoter. J Bacteriol 183: 2842–2851. Takeda H, Yamada A, Miyauchi K, Masai E, Fukuda M (2004) Characterization of transcriptional regulatory genes for biphenyl degradation in Rhodococcus sp. strain RHA1. J Bacteriol 186: 2134–2146. Thanbichler M, Iniesta AA, Shapiro L (2007) A comprehensive set of plasmids for vanillate- and xyloseinducible gene expression in Caulobacter crescentus. Nucleic Acids Res 35: e137. Towe S, Leelakriangsak M, Kobayashi K, Van Duy N, Hecker M, Zuber P, Antelmann H (2007) The MarR-type repressor MhqR (YkvE) regulates multiple dioxygenases/glyoxalases and an azoreductase which confer resistance to 2-methylhydroquinone and catechol in Bacillus subtilis. Mol Microbiol 66: 40–54. Tropel D, Bahler A, Globig K, van der Meer JR (2004) Design of new promoters and of a dual-bioreporter based on cross-activation by the two regulatory proteins XylR and HbpR. Environ Microbiol 6: 1186–1196. Tucker JB, Zilinskas RA (2006) The promise and perils of synthetic biology. New Atlantis 12: 25–45. Urata M, Miyakoshi M, Kai S, Maeda K, Habe H, Omori T, Yamane H, Nojiri H (2004) Transcriptional regulation of the ant operon, encoding twocomponent anthranilate 1,2-dioxygenase, on the carbazole-degradative plasmid pCAR1 of Pseudomonas resinovorans strain CA10. J Bacteriol 186: 6815–6823. Velazquez F, de Lorenzo V, Valls M (2006) The m-xylene biodegradation capacity of Pseudomonas putida mt-2 is submitted to adaptation to abiotic stresses: evidence from expression profiling of xyl genes. Environ Microbiol 8: 591–602. Velazquez F, Parro V, de Lorenzo V (2005) Inferring the genetic network of m-xylene metabolism through expression profiling of the xyl genes of Pseudomonas putida mt-2. Mol Microbiol 57: 1557–1569.

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Vesely M, Knoppova M, Nesvera J, Patek M (2007) Analysis of catRABC operon for catechol degradation from phenol-degrading Rhodococcus erythropolis. Appl Microbiol Biotechnol 76: 159–168. Vitale E, Milani A, Renzi F, Galli E, Rescalli E, de Lorenzo V, Bertoni G (2008) Transcriptional wiring of the TOL plasmid regulatory network to its host

involves the submission of the sigma-promoter Pu to the response regulator PprA. Mol Microbiol 69: 698–713. Wall ME, Hlavacek WS, Savageau MA (2004) Design of gene circuits: lessons from bacteria. Nat Rev Genet 5: 34–42.

41 Emerging Systems and Synthetic Biology Approaches to Hydrocarbon Biotechnology V. de Lorenzo* . S. Fraile . J. I. Jime´nez Systems Biology Program, Centro Nacional de Biotecnologı´a-CSIC, Campus de Cantoblanco, Madrid, Spain *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1412 2 Describing Complexity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1414 3 Systems Biology of One Bacterium at a Time . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1417 4 From Genomes to Catalysts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1420 5 Systems Biology of In Situ Bioremediation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1422 6 The Catalytic Gene Landscape . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1423 7 Syntrophic and Ectopic Metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1426 8 Predicting Catalysis and Biodegradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1428 9 Outlook: From Systems to Synthetic Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1430

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_97, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Systems Biology is a conceptual frame for studying living systems that departs from the extreme reductionism of traditional Molecular Biology and pursues the quantitative understanding of complete biological entities rather than the mere comprehension of their parts. The development of the field stems from the popularization of high-throughput DNA sequencing technologies that have allowed the complete determination and archiving of the whole genetic complement of a number of individual species from microorganisms, and even complete ecosystems. This has then been followed by a plethora of omics disciplines (genomics, proteomics, metabolomics) that endow Biotechnologists upfront with wealth of information on any biological catalyst of interest. This article examines questions at stake in the application of Systems Biology to microbial biotransformations of petroleum components, in particular, the type of abstractions needed for modeling the scenarios where microorganisms meet organic molecules. Concepts such as epi-metabolome, pan-enzymes, and ectopic metabolism are paramount to comprehend microbial activities in sites where mixtures of chemical structures are exposed to multispecies metabolic networks. The ensuing process is constrained by the abiotic characteristics of the locations, as physico-chemical conditions determine the dynamic interplay between the contaminants and the biological catalysts. The availability of genes, genomes, and metagenomes of biodegradative microorganisms make it possible to model and predict the fate of chemicals through the global microbial metabolism. Moreover, the field is capitalizing quickly on the new field of Synthetic Biology (SB) in view of the possibilities of designing superior biocatalysts for biodegradation and biotransformations of desired chemicals.

1

Introduction

There are a large number of scenarios – natural and man-made – where microorganisms are either directly exposed to petroleum compounds or to organic chemicals derived from petroleum. In the simplest case, one given natural or recombinant strain endowed by a certain catalytic activity faces one single chemical under the controlled conditions of a reactor for its biotransformation in an added value product or for its biodegradation into CO2 and H2O. In the other extreme, we have cases of complex, multi-strain microbial communities being exposed to mixtures of chemicals with diverse structures in an open environment with diverse physicochemical conditions that undergo changes on which we have little or no control (Semple et al., 2007). Both extremes define the boundaries of Petroleum Biotechnology (and even Chemical Biotechnology at large), but we find a quite different situation in each of the two edges. The use of microorganisms as catalysts for biotransformations of chemicals in a bioreactor has been a thriving field or research and application for many decades (Schmid et al., 2001). Much before the onset of genetic engineering, a considerable number of natural microbial isolates were used by organic chemists for adding value to otherwise low-cost substrates. This was followed in most cases by identification of the biological activities underlaying such biotransformations and the cloning of the corresponding genes. As soon as the genes of interest started to be cloned, the possibilities of exploiting naturally occurring biotransformations for process engineering started to boom owing to the many possibilities offered by modern DNA technology (Heinemann and Panke, 2006). Microorganisms then became cell factories for either production of valuable chemicals or for destruction of toxic waste. The panoply of tools to this end includes (1) systems for overexpression of cloned genes, (2) mobilization of the desired catalytic activities into microbial hosts specialized in enduring the harsh working

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conditions of an industrial bioreactor, (3) random or directed mutagenesis of genes or groups of genes for a superior catalytic performance or for changing substrate profiles, (4) mining of new enzymes in environmental metagenomes, (5) forward design of proteins for either improving existing properties or for creation of altogether new ones, and (6) sophisticated process engineering for optimal biotransformation yields. Many of these successes have applied directly to biotransformations of interest to the oil industry (see Van Hamme et al., 2003 for a review). One remarkable case involved the genetic engineering of bacterial catalysts for enhanced desulfurization of dibenzothiophene (Calzada et al., 2008; Gallardo et al., 1997). These conceptual and material tools, now under the umbrella of Green Chemistry, are growingly considered as tokens of the sustainability of the chemical industry of the future. Yet, until very recently, most breakthroughs in the field of industrial biotransformations have originated in trial-and-error approaches resulting better strains, better genes, better proteins, better growth media, etc. In fact, the part of petroleum biotechnology related to transformations of industrial interest in controlled reactors has benefited in nearly real time from the progress made first in Microbial Genetics and then in Molecular Biology. Furthermore, the complexity of the working scenarios in reactors is considerable but still limited and tractable, since many conditions can be fixed at the will of the operator: water saturation, media composition, O2 contents, temperature, C and N sources, electron donors/acceptors, etc. The situation is far less rosy if one considers the other extreme of the microbe-petroleum scenario. In this case, complex microbial consortia with thousands of different species interact in open aquatic and terrestrial environments with a large number of chemical species. These instances typically include, e.g., petroleum spills in the sea or soil polluted with industrial chemical waste. In these cases, the catalytic activities of the biological party and their outcome on the chemicals are determined not only by the genes encoded in the genome, but also by a much larger number of variables. Apart from the multispecies nature of this type of settings, the changeable composition and the diverse physico-chemical conditions of the place at stake make a huge difference. The bioavailability of the chemicals, the water tension, oxygen, temperature, UV light, and other abiotic conditions determine the type of microorganisms capable of colonizing the niche (Semple et al., 2007). These can in turn have or not catalytic genes of interest and can express them or not. Initial conditions can also change as microbial activities alter the chemical landscape of the place, which consecutively can modify the structure of the microbial community, and so on (de Lorenzo, 2008). The number of variables is so high that molecular information on catalytic genes of specific strains (which is so useful for controlled biotransformations) often becomes intractable for understanding such an in situ catalysis. Needless to say that the problem is complexity. In this context, this article addresses the management and exploitation of microbiological complexity for environmental biocatalysis with the tools of Systems Biology. Although much abused, the term still expresses an agenda of quantitative study of complex interactions in biological systems using an integrative perspective – instead of reductionist one – to study them (Alon, 2006; de Lorenzo, 2008; Fredrickson et al., 2008; Trigo et al., 2008). One of the key goals of Systems Biology is to unveil emergent properties embodied in the inner organization of complete biological systems. To this end, any System Biology approach is organized in three stages, i.e., description of the system (for which all types of omics data thereof are to be considered), deconstruction of the system in its components, and, eventually, reconstruction of the system with the same or with other properties. Note that the term deconstruction has two different meanings, both of them incorporated to the Systems Biology jargon. First, it means the stepwise dismantlement of the components of an object for reuse, recycling, or management. This is the intuitive meaning of the term, as it

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evokes the breakdown of, e.g., a piece of furniture or a radio into singular, basic pieces. But deconstruction also denotes the uncovering of an implicit or hidden significance in a text or in an object, which is not apparent from a superficial description. This meaning (less familiar to biologists) is certainly part of the agenda of Systems Biology. Finally, reconstruction of a system is the ultimate proof of understanding, echoing the celebrated remark by the 1965 Nobel Prize winner in Physics Richard Feynman ‘‘what I cannot create, I do not understand.’’ Synthetic Biology (SB) takes this last aspect further to the point of proposing the design of nonnatural biological systems following a rational blueprint (Arkin, 2008; Canton et al., 2008), which results in properties a´ la carte for biotechnological applications. Synthetic Biology and its projection into petroleum Biotechnology will be addressed later in this article. The following sections tackle the challenge of quantifying complexity, the applications of Systems Biology to singular catalysts, and some views of Bioremediation and environmental catalysis from a Systems perspective. Finally, we will examine how Synthetic Biology can provide a fresh start to the challenge of designing not only superior catalysts for industrial biotransformations, but also safe and efficacious microbial catalysis for in situ removal of toxic waste.

2

Describing Complexity

It has been argued before that Physics is the one and only Science and that any other body of knowledge becomes more scientific only when it adopts the rigorous descriptive language of Physics and its analytical and predictive powers (Wolpert, 1998). There is some echo of this in the fact that modern Molecular Biology was founded by physicists rather than by biologists or naturalists (Schro¨dinger, 1945). In that case, however, the greatest contribution of Physics to the development of Biology was the adoption of formal logics, abstractions, and extreme reductionism for understanding biological phenomena. The basic idea is that one can always break down a complex system into smaller subsystems with a limited number of components to the point of making them amenable to thorough logic analyses. The dividends of such an approach need no further justification, as reductionism is the conceptual frame that has allowed the breathtaking development of Molecular Biology and Biotechnology since the late 1940s. However, in live organisms, the whole is more than the sum of the parts, and some of the properties of the system cannot be obtained from the properties of its individual components, so requiring the study of the biological object as a whole. We must clarify now the meaning of complexity. In colloquial language, the term is used to qualify something with many parts in intricate arrangements. But in rigorous scientific language, complexity is the quality of one system composed of a number of elements and a number of relationships among them, which can be differentiated from those with other elements outside the relational body. This means that complexity is an operative, relative term rather than an absolute feature of a system. A simple way of describing complexity is, therefore, to identify the elements of a system and to recognize and categorize their interactions. There are several formal ways of recounting the complexity of a system. But by far the most popular in Systems Biology is the language of networks and network theory. This field was pioneered by Albert-La´szlo´ Barabasi (Barabasi and Bonabeau, 2003; Jeong et al., 2000) and thoroughly applied to Biological systems by Uri Alon (Alon, 2006, 2007). The first studies of the properties of biological networks (e.g., metabolic networks, protein interaction networks, genetic control networks) revealed new facets of living systems. One of the main ideas to come out of this research was that the topology of those biological networks was not random but had a

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typical structure called scale-free (Jeong et al., 2000). In these networks the distribution of connectivity is not homogeneous (> Fig. 1a) but follows a power law: there are a few highly connected nodes (hubs) and the rest has low connectivity, in contrast with random networks where the connectivity follows a Poisson distribution. Scale-free networks have two main properties: the pathway between any two nodes is always short because the hubs act as shortcuts, and they are very tolerant against random perturbations (elimination of components) because there are always alternative pathways through the hubs. Further research has unveiled distinct motifs in biological networks (Alon, 2007) that endow the systems under scrutiny with global properties that could not be deduced otherwise from singular components. The downside of this straight representation of networks is the difficulty to enter directional actions and transmit signals (increase/decrease, activate/repress, etc.) between the components of a network, as is the case in biological systems. In these instances, the network structure is often the basis of deterministic models that can be represented as a whole of differential equations that establish the relationships between the parts (Kaern et al., 2003). This can be done properly only when many –if not all – of the parameters that rule interactions in the system are known. However, the biochemical mechanisms underlying the interactions are often scarce or unknown, which prevents the formulation of detailed models. Furthermore, information on kinetic parameters and molecular concentrations is typically limited or absent. Therefore, traditional methods for numerical analysis are difficult to apply. A convenient way to represent actions in a network when either there are not enough data or when the data are available in heterogeneous formats is the integration of different input signals in Boolean, binary fashion that can be implemented through logic gates (Silva-Rocha and de Lorenzo, 2008). Such gates perform operations on one or more inputs and produce each time a single logic output (> Fig. 1b). Since the output is also a logic-level value, an output of one logic gate can connect to the input of one or more other logic gates. The logic thereby performed is thus adequate for the functioning of digital circuits. Logic gates are typically implemented electronically using diodes or transistors, but can they also be shaped with biological components (Silva-Rocha and de Lorenzo, 2008). Although binary logic circuits are based on functions with just two possible states (0 or 1), existing biological systems typically display continuous values for the input/output functions. Fortunately, there are stratagems for matching biological data, even if they are not quantitative, to binary representations such as high/ low, active/inactive, etc. Often such simple formalisms reveal aspects of regulatory systems that are otherwise hidden in a merely descriptive narrative. An approach that is capable of handling many of the above problems is based on a class of piecewise-linear (PL) differential equation models originally proposed by Glass and Kauffman (1973). The state variables in the PL models correspond to the concentrations of proteins encoded by the genes in the network, while the differential equations represent the interactions arising from the regulatory influence of some proteins on the synthesis and degradation of others. The regulatory interactions are modeled by means of step functions, giving rise to PL structures of differential equations. PL models have been successfully used for the study of several prokaryotic and eukaryotic regulatory networks and holds a considerable promise to examine cases where not all the elements of a given organisms of system are known (de Jong et al., 2003). Useful as they are, deterministic models of the type just mentioned above are not good enough for describing biological behavior at the level of single cells, where basically all events are stochastic (Shahrezaei and Swain, 2008). Phenotypes do vary across isogenic populations and in individual cells over time. Even in an isogenic population, every cell is unique, specially in the expression of their genes. Stochasticity is enhanced in cases where just a few of the molecules are

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. Figure 1 Formalization of relational objects in Systems Biology. (a) Networks. The simplest way of describing a complex system is to represent it as network in which a number of nodes are connected by a number of edges. The topology of the network can be expressed in part as the relation between the connectivity of each of node, k, and the number of nodes with a given number of connections p(k). In scale-free networks, the connectivity follows a power law distribution, p(k) = k-g. In virtually all metabolic networks explored thus far, the exponent g has a value between 2 and 3. Note a small number of very connected nodes (hubs) and many nodes with very few connections. (b) Logic gates. Actions between components of a network can often be formalized through the use of logic gates. Typically, such gates perform operations on two inputs (each of them having a 0/1 value) and produce each time a single logic output 0/1. Since the output is also a logic-level value, an output of one logic gate can connect to the input of one or more other logic gates. Logic gates are typically implemented electronically using diodes or transistors but can they also be constructed using inter alia promoters and regulators. The symbols of the three most frequent gates (AND, OR, NOR) and the tables of truth associated to each of them are shown.

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involved (e.g., regulatory events), since low numbers make individual reaction events more significant. Any phenotype measured from a population of cells or from a single cell at different times will not have a unique value, but a collection of values. Stochastic events are generated by the dynamics of the system from the random timing of individual reactions (Elowitz et al., 2002). Noise is therefore an empirical measure of stochasticity (Pedraza and van Oudenaarden, 2005). The growing availability of technologies to monitor (mostly regulatory) events in single cells allows the processing experimental data taking into account stochasticity both at the modeling stage and in the design of the identification algorithms (Chabot et al., 2007; Cinquemani et al., 2008). Yet, there is still a considerable challenge to translate single-cell stochastic behavior into population performance. The projection of these novel system analysis tools to oil-related microbial processes is still in its infancy but they will certainly make a difference in the modeling and actual applications of designer catalysts that lay ahead of us.

3

Systems Biology of One Bacterium at a Time

One useful start to employ Systems Biology approaches to oil microbiology and biotechnology is the perusal of the singular genomes of bacteria that display some type of biological activity on petroleum components. One of the first attempts in this direction regards the complete sequence of the TOL plasmid pWW0 of Pseudomonas putida mt-2 for catabolism of toluene and m-xylene (Greated et al., 2002). The xyl genes borne by the TOL system are one of the best studied single instance of a bacterial system able to degrade aromatic hydrocarbons, including its genetics, regulation, and biochemistry. The sequences of many other catabolic plasmids encoding genes for degradation of recalcitrant or xenobiotic chemicals have been reported and cataloged since (Leplae et al., 2006). Following pWW0, the genomic sequence of P. putida KT2440, the natural host of the TOL plasmid was completed as well (Nelson et al., 2002). This strain of Pseudomonas, one of the best studied paradigms of bacteria of biotechnological interest for this type of application, is the subject of other chapters of this book (> Chapter 22, Vol. 2, Part 5) and will not be examined in detail here. In any case, P. putida KT2440 remains to this day as one favorite experimental system to test the power of Systems Biology when applied to a bacterial catalyst (Nogales et al., 2008; Puchalka et al., 2008). Much work on this strain is currently in progress and several industrial and environmental applications are likely to be added soon to those currently available. A good share of public databases listing sequenced genomes (see for instance, http://www.ncbi.nlm.nih.gov/Genomes) consists of strains known to have catalytic properties for biodegradation or biotransformations of various types of chemicals. Although a considerable number of singular strains have dedicated chapters in this book, we comment below a small selection of those that we find particularly interesting from a Systems Biology point of view. The 9.73 Mbp multi-replicon genome of Burkholderia xenovorans LB400 (formerly called Pseudomonas sp. LB400 and B. cepacia LB400) was reported in 2006 (Chain et al., 2006). This strain was known for a long time to be one of the best aerobic degraders of polychlorinated biphenyls (PCBs) (Perez-Pantoja et al., 2008). Analysis of the genome revealed the presence of not only 11 central aromatic pathways, but also 20 additional peripheral routes for other aromatics (including redundant benzoate degradation routes) and formaldehyde oxidation. The apparently stable coexistence of three replicons is intriguing and says about the genomic plasticity of this strain. Many other genetic factors are associated with in vivo survival and intercellular interactions, what surely widens the niche breadth of this bacterium. A gram-positive

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counterpart of B. xenovorans LB400 is Rhodococcus sp. RHA1 (McLeod et al., 2006). This soil actinomycete was sequenced also in 2006 and its genome size and catabolic versatility rival those of LB400. Rhodococcus sp. RHA1 is a potent PCB-degrader with one of the most complex bacterial genomes known, as it includes a linear chromosome and three linear plasmids. Its DNA is exceptionally rich in coding oxygenases, many of which occur in the >30 pathways predicted to degrade aromatic compounds. RHA1 and LB400 thus appear to demonstrate that similar biodegradative capacities can nest within very different bacterial types. Cupriavidus necator JMP134 (formerly and consecutively called Alcaligenes eutrophus, Ralstonia eutropha, and Wautersia eutropha) is another bacterium with amazing biodegradative abilities that has been recently sequenced (Perez-Pantoja et al., 2008). The strain was isolated from an Australian soil by its ability to grow on 2,4-dichlorophenoxyacetate (2,4D), but later studies showed that it grew also on 4-methyl-2-chlorophenoxyacetate and 3-chlorobenzoate (3CB) The determinants for 2,4D and 3CB degradation (the tfd genes) have been known for a long time to reside in the catabolic plasmid pJP4. However, only the reconstruction made possible with the complete genomic sequence has exposed the extensive metabolic power that this bacterium is endowed with (Perez-Pantoja et al., 2008). More than 60 aromatic chemicals serve as a sole carbon and energy source for this strain, strongly correlating with catabolic abilities predicted from genomic data. A broad spectrum of peripheral reactions seems to channel substituted aromatics into not less than ten central ring-cleavage pathways. Gene redundancy seems to play a significant role in the catabolic potential of this bacterium, posing the question on how this strain avoids metabolic noise. These qualities make the genome of C. necator JMP134 a virtually complete toolbox of catabolic genes and regulators for engineering biocatalysts aimed at degradation of aromatic chemicals. The genomic sequence of Alcanivorax borkumensis SK2 was the first example of a marine hydrocarbonoclastic microorganism (Schneiker et al., 2006). The most interesting aspect of this aerobic gram-negative g-proteobacterium is not that it degrades hydrocarbons – many marine and other bacteria do this – but that it lives exclusively on alkanes, ignoring other standard carbon sources like sugars and amino acids. In addition, A. borkumensis does not consume the light aromatic fractions that many environmental bacteria are able to metabolize. Because of these properties, it comes as no surprise that boomings of Alcanivorax are linked to the removal of hydrocarbons in fertilized, oil-contaminated seawater. The repertoire of diverse alkane hydroxylases (which initiate degradation of branched alkanes very efficiently) make these bacteria major players in the spontaneous biological cleanup of oil-contaminated environments. Given the reduced number of external nutrients that A. borkumensis SK2 uses, it is likely that most of the regulatory processes are related to global balancing of carbon, nitrogen, and phosphorus in the context of enduring environmental stresses, rather than to a response to specific small signal molecules (Reva et al., 2008). On this basis, one could argue that the key to improving oil biodegradation in situ is to focus on the nutritional balance of indigenous marine bacteria by careful management of the carbon to nitrogen to phosphorous ratios. This simple notion (rather than inoculation of oil-degrading superbugs) should guide bioremediation strategies in the future for these types of environmental problems (de Lorenzo, 2006). The genome of Shewanella oneidensis MR1 was reported in 2002 as an important model organism for bioremediation because of its diverse respiratory capabilities, conferred in part by multicomponent, branched electron transport systems (Heidelberg et al., 2002). Specifically, the strain received much attention because of its ability to reduce uranium and chromium. It was soon realized that the versatility of Shewanella spp. extended to other nonstandard metal electron acceptors, such as iodate, technetium, neptunium, plutonium, selenite, tellurite, and

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vanadate, and even the reduction of nitroaromatic compounds by some strains. These diverse metabolic capabilities provide shewanellae with a considerable potential for the remediation of environments that are contaminated with radionuclides (Fredrickson et al., 2008). One fascinating application of this bacterium in recent years comes from the fact that reduction of soluble Pd(II) by cells of S. oneidensis MR1 and of an autoaggregating mutant (COAG) results in precipitation of palladium Pd(0) nanoparticles on the cell wall and inside the periplasmic space (bioPd). The bioPd(0) nanoparticles thus obtained have the ability to reductively dehalogenate PCB congeners and lindane in aqueous and sediment matrices (De Windt et al., 2005; Mertens et al., 2007). This opens interesting possibilities of engineering bio-inorganic materials for bioremediation in which biological and abiotic catalyses merge for the sake of degrading a target contaminant. Rhodopseudomonas palustris is a purple photosynthetic bacterium that ranks among the most metabolically versatile microbes known (Larimer et al., 2004). Specifically, it grows by any one of the four modes of metabolism that support life: photoautotrophic or photosynthetic (energy from light and carbon from carbon dioxide), photoheterotrophic (energy from light and carbon from organic compounds), chemoheterotrophic (carbon and energy from organic compounds), and chemoautotrophic (energy from inorganic compounds and carbon from carbon dioxide). R. palustris enjoys exceptional flexibility within each of these modes of metabolism. It grows with or without oxygen and uses many alternative forms of inorganic electron donors, carbon and nitrogen. It degrades plant biomass and chlorinated pollutants and it generates hydrogen as a product of nitrogen fixation. The sequence of strain CGA009 of this species was reported in 2003 and found to encode 3 nitrogenases, 5 benzene ring-cleavage pathways, and 4 light harvesting systems. In addition, R. palustris bears 63 signal transduction histidine kinases and 79 response regulator receiver domains. This makes P. palustris an organism of choice to probe how the web of metabolic reactions that operates in single cell adjusts itself in response to changes in light, carbon, nitrogen, and electron sources. The most remarkable application of this bacterium thus far has been the refactoring of its metabolism for achieving a high production of H2 (Rey et al., 2007), but given its metabolic plasticity it will also be a good choice as a chassis for implantation of engineered biodegradation pathways. Desulfovibrio vulgaris is archetypical example of anaerobic, sulfite-reducing bacteria that contribute to bioremediation of toxic metal ions. This is because their metabolism increases the pH, causing toxic divalent metal ions like copper, nickel, and cadmium to precipitate as metal sulfides in acidic aquatic environments (e.g., mine drainage). These bacteria can also deliver electrons directly to oxidized toxic metal ions, including uranium (VI), technetium (VII), and chromium (VI), converting these into less-soluble, reduced forms, which are less bioavailable. Metal reduction mediated by this type of bacteria thus represents a potentially useful mechanism for the bioremediation of metal ion contaminants from anaerobic sediments. Publication of the complete sequence of the Hildenborough strain of D. vulgaris (Heidelberg et al., 2004) allowed the identification and initial characterization of the bacterium’s complex, periplasmic cytochrome network and cytoplasmic sulfate reduction capabilities. The sequence has also allowed in-depth characterization of cell-wide responses to low-oxygen exposure (Mukhopadhyay et al., 2007) and some mechanisms for biocorrosion – an undesirable downside of this type of microorganisms. In any case, the number of anaerobic or facultatively anaerobic strains that have been sequenced because of their biocatalytic activities on pollutants is still small. The denitrifying strain Azoarcus sp. EbN1 (just renamed Aromatoleum aromaticum EbN1 [Wohlbrand et al., 2008]) is unique in anaerobically degrading alkylbenzenes via different pathways, which converge at benzoyl-coenzyme A. Analysis of its genomic sequence

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(Rabus et al., 2005) shows that this strain contains ten additional anaerobic and four aerobic aromatic degradation pathways, which seem to coexist and become alternatively activated in response to the substrates and the environmental O2 tension. This is reflected in the presence of paralogous gene clusters (e.g., for anaerobic phenylacetate oxidation) and high sequence similarities to orthologs from other strains (e.g., for anaerobic phenol metabolism) along with multiple respiratory complexes. Anaerobic processes are extremely attractive for in situ bioremediation of many types of pollutants in subsurface environments. For some bacteria, the target contaminants serve as electron acceptors rather than electron donors in reactions that occur in anoxic sites. Geobacter species (several strains of which have been sequenced [Butler et al., 2007]) can oxidize a large number of organic compounds concomitantly with the reduction of Fe (III) to Fe (II). Geobacter can also use metals as electron acceptors for respiration. Although the reduction of metals does not destroy them, it often changes their solubility and makes them less toxic. For instance, some Geobacter strains can reduce the soluble, oxidized form of uranium, U (VI), to the insoluble form, U (IV). This precipitates the uranium from contaminated groundwater, thereby preventing its further spread. The opportunities of improving these processes through genetic engineering are developing very rapidly (Izallalen et al., 2008). A different type of anaerobic process that can be capitalized in environmental catalysis is reductive dechlorination, in which microorganisms remove chlorine atoms from chlorinated contaminants (e.g., PCBs and the like), by using these chemicals as electron acceptors in respiration. Various strains of Dehalococcoides, one genus that seems to be equipped the best for catalyzing these reactions in the subsurface, have been sequenced ([Seshadri et al., 2005]; see also updates in http://www.ncbi.nlm.nih.gov/Genomes). But how the genomic information of Geobacter or Dehalococcoides strains could inspire new bioremediation schemes? Current biostimulation strategies contemplate the addition of nutrients and electron donors or acceptors to trigger activities involved in the bioremediation setup (N’Guessan et al., 2008) or even bioaugmentation with natural or genetically improved strains (see below). In fact, inoculation of target sites with Geobacter seems to be one of the few bioaugmentation success stories (Lovley, 2003). Perhaps under anoxic conditions, the grazing of the extra biomass by resident protozoa mentioned below may not be that significant.

4

From Genomes to Catalysts

The onset of Systems Biology (and more recently, Synthetic Biology) has relaunched the objective of creating in the laboratory designer microorganisms with superior catalytic abilities on recalcitrant pollutants (Brenner et al., 2008; Cases and de Lorenzo, 2005; de Lorenzo et al., 2006; de Lorenzo and Danchin, 2008). Metabolic engineering is a whole research field by itself and will not be covered in detail here. However, some methodological and conceptual advances are worth to mention in connection with Biocatalysis of oil-related chemicals. One relevant aspect is the rational change of substrate specificity of certain enzymes for overcoming bottlenecks in any given metabolic pathway. It is growingly believed that extant enzymes with high specificity and activity have evolved from ancestral enzymes with promiscuous functions (Afriat et al., 2006; Peisajovich and Tawfik, 2007). This process is highly dependent on the ability of proteins to alter their capacities with a small number of amino acid substitutions. Identification of such plasticity residues (which are not necessarily coincident with those that endow specificity) is therefore of essence to set up experimental

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evolution systems to allow emergence of new enzyme functions (Yoshikuni et al., 2006). A second development is the design of altogether new reaction centers on suitable protein scaffolds that do not have per se any enzymatic activity (Dwyer and Hellinga, 2004). Although this notion, pioneered by H. Hellinga, has been recently subjected to heavy criticism (Hayden, 2008), it is still one of the most promising approaches to generate new enzymes. Finally, there is issue of assembling nonnatural operon-like multi-gene metabolic pathways with a balanced expression of the corresponding products. This general problem has been met by the use of libraries of tunable intergenic regions that act as posttranscriptional control elements and facilitate the screening for the most suitable relative expression levels (Pfleger et al., 2006). Promoter libraries with defined levels of expression are also available to the metabolic engineer for tuning the presence of the implanted pathways to the general well-being of the host cell. The next challenge for designing bacterial catalysts a´ la carte is the implantation of a new function or a complete pathway within the existing metabolic network of the host, as the background metabolic complexity may not fit well with such an implant (Alper and Stephanopoulos, 2008). There are a number of computational tools to examine this question beforehand, some of which are worth to highlight. For instance, the Optkock framework (Burgard et al., 2003) is a system originally developed in Escherichia coli for overproduction of chemicals, but the concept can be extended to many other bacteria as well. Starting with a genome-based metabolic model, Optknock matches production of a desired product to the stoichiometric drain of growth resources (i.e., carbon, redox potential, and energy). In this way, the platform proposes deletion strategies for eliminating competing reaction pathways as well as other mechanisms of compensating for the removed functionalities. The system has found one very useful application in the design of Geobacter strains with increased respiration rates (Izallalen et al., 2008). Another platform called Optstrain uses a database of bioreactions to elucidate the set(s) of functionalities that are to be added to a given host to achieve formation of a desired product (Pharkya et al., 2004). The Monte Carlo algorithm called DESHARKY (Rodrigo et al., 2008) finds a metabolic pathway from a target compound by exploring a database of enzymatic reactions. This system outputs a biochemical route to the host metabolism together with its impact in the cellular context by using mathematical models of the cell resources and metabolism. Most modeling tools of this sort are directed to generation of products (Tyo et al., 2007) rather than to their biodegradation. Finally, the many changes required for designing optimally microorganisms for biocatalysis can benefit from the growing possibility to synthesize whole genomes for desired biotechnological applications. While synthesizing DNA molecules of the size of a bacterial genome is becoming a realistic option (Gibson et al., 2008), it is also true that the difficulties of achieving the desired results become greater as increasingly complex sets of genes are brought together. Apart from metabolic engineering, design of bacteria for the environment touches upon other constraints that limit the performance of any molecular machinery within the overcrowded volume of a bacterium. One emerging feature of the intracellular organization of bacteria is that most polypeptides assemble into multi-protein structures (Norris et al., 2007; Su et al., 2007) so that most proteins are not singular objects let loose in the cytoplasm, but pieces of multipart, dynamic structures that assemble through a constellation of specific interactions (Norris et al., 2007). This means that each protein needs to find a suitable intracellular niche for optimally performing its function. Polypeptides unable to fit within such assemblies might be rendered not functional and eventually rejected through a simple Darwinian mechanism. Implantation of new proteins in a cell can thus be severely counterselected (Lercher and Pal, 2007; Sorek et al., 2007). How to escape this constraint? One

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possibility is to let the implanted pathway to evolve in vivo until it finds the most optimal configuration of gene doses and expression levels, which are compatible with the physiology of the host thereby facilitating their nesting in the recipient protein network (see for instance, Camps et al. [2003] as a possible technique to this end). Alternatively, the implanted proteins and functions could be designed for having a minimal dependence on the biological context of the recipient. The quality of behaving in a context-free fashion is designed as orthogonality,1 echoing equivalent properties in computing science. The pursuit of orthogonal metabolic modules might thus provide a new perspective in metabolic engineering of catalytic bacteria.

5

Systems Biology of In Situ Bioremediation

Bioremediation is mostly about intervention aimed at alleviating pollution. In this sense, the field is not to be confounded with biodegradation, which tackles the biological bases of the (mostly bacterial) metabolism of unusual and/or recalcitrant compounds. Depending on the degree of such intervention, bioremediation is generally considered to include natural attenuation (little or no human action), biostimulation (addition of nutrients and electron donors/acceptors to promote the growth or metabolism of certain microorganisms), or bioaugmentation, the deliberate addition of natural or engineered microorganisms with the desired catalytic capabilities (El Fantroussi and Agathos, 2005; Singer et al., 2005). Bioremediation as a technology to get rid of undesirable waste has been going on in history much before the onset of modern Biology. Black wells of mediaeval Italian cities, or nineteenthcentury waste treatment plants can be considered early instances of ex situ bioremediation operations. Yet, these primitive setups operated on a black box principle, the functioning of which remained mysterious until more recent times: deliberate use of microbial activities to detoxify environmental contamination was first applied to hydrocarbon-contaminated groundwater systems in the early 1970s. On the other hand, the modern scientific field of biodegradation was initiated in the mid-1960s in the USA by I. C. Gunsalus, D. Gibson, and N. Ornston, and by H. Knackmuss in Germany, with their studies on the metabolism of organic compounds by Pseudomonas-type strains. Since then, a large number of gram-positive and gram-negative bacteria, as well as fungi and archaea have been isolated, which are able to grow or to co-metabolize many compounds that are typical environmental pollutants (Diaz, 2004; Pieper et al., 2004). The popularization of recombinant DNA technology since the mid-1980s and the more recent possibility to fast sequence large sequences of DNA opens the possibility to convert the basic knowledge derived from such fundamental data on the genetics and biochemistry of biodegradation into real bioremediation actions. One distinct angle of the problem is whether advanced metabolic engineering can result in microorganisms with superior catalytic activities which – one introduced in a given polluted site – accelerate the removal of toxic waste. The attractive idea (so popular through the 1990s [Pieper and Reineke, 2000; Timmis and Pieper, 1999]) that one can isolate bacteria with interesting biodegradative activities on recalcitrant compounds improve them genetically and put them back into the environment as superior bioremediation agents have failed to meet the expectations (Cases and de Lorenzo, 2005). Yet, along the way interesting features have been revealed, which are of essence for

1

A is orthogonal to B, if A does not influence B. Orthogonality guarantees that changes made in a component of a system neither creates nor propagates side effects to other components of the system.

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future attempts in Environmental Biotechnology. First, most biodegradative microorganisms studied so far have been isolated in enrichment cultures. Once singular strains are isolated, conventional biodegradation studies typically focus on the interplay between one bacterium and one chemical species. The logic of enrichment links biodegradation to a faster buildup of biomass, so those isolated are the ones, which convert more efficiently the C backbone of the pollutant into metabolic intermediates, not to CO2 and water (Janssen et al., 2005). While the best bioremediation agents would be those able to enter maximum catalytic activity with a minimum of biomass, the onset of fast growers brings about a niche for protozoa (El Fantroussi and Agathos, 2005; Lueders et al., 2004), which end up destroying the very catalysts. In fact, the use of stable isotope probing (SIP) has proven without a doubt that the members of natural bacterial consortia that degrade given chemicals in situ are not those that may appear in enrichment cultures (Wackett, 2004). A second aspect of the divide between biodegradation and bioremediation is that enrichment cultures typically result in single strains able to execute a complete biodegradation pathway. Yet, it is possible that such strains do not occur naturally, but they originate in the gene transfer events and the strong evolutionary selection associated to long enrichment procedures starting with a complex environmental sample. By the same token, the combined activities of various strains in a consortium may afford degradation of chemicals, which cannot be performed in by single strains, and thus cannot be isolated through enrichment. Also, polluted sites most often occur contaminated by numerous chemicals, so that a whole of chemical structures is exposed to the catalytic abilities of multiple strains. How to identify who is who and who makes what on whom in such a multi-actor scenario? Although new enrichment strategies and growth-independent activity screening have been proposed for selection of biodegradative strains and communities the experience generally discourages the use of fast growers as bioremediation agents (see above). Finally, enrichment procedures are blind to non-culturable microorganisms, what constitutes the larger fraction of the microbiota and surely form the largest repository of catalytic activities (Ferrer et al., 2005). To make things more challenging, the factors at play in bioremediation scenarios include more elements than just the biological catalysts and the contaminants discussed above. Their dynamic interactions occurs in concrete abiotic settings, which are defined by a whole of physico-chemical conditions: O2 tension, electron acceptors, water, temperature, granulation, and others, many of which change over time and the course of the catalysis (Eyers et al., 2006; Katsivela et al., 2005; Wenderoth et al., 2003). Such abiotic conditions determine the species composition of the endogenous microbial communities as much (or more) than the availability of given chemical species as carbon and energy source. Bioremediation is thus a case of multiscale complexity, which is not amenable to the typically reductionist approaches (e.g., one compound, one strain, one pathway) that have dominated studies on Biodegradation.

6

The Catalytic Gene Landscape

A number of abstractions are useful for representing the part of the microbial community present in a given niche that is relevant for in situ bioremediation. The first consideration is that what matters in bioremediation is the presence and performance of the catabolic activities available in the site, regardless of the particular species that carry them (de Lorenzo, 2008). The second issue is the division of the metabolome of any given microorganism in three categories with distinct diffusion abilities: (1) an intrinsically nondiffusible pool of metabolites,

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which never make it outside the cells; (2) a diffusible metabolome composed of molecules that can occasionally be secreted depending on the catalytic rate of the bacteria, which produce them; and (3) a peripheral metabolome (epi-metabolome) formed by the pool of compounds, which are transformed so slowly that they can diffuse out the cells between one step of a metabolic pathway and the next one – or are actively secreted because of its toxicity (de Lorenzo, 2008). Such a free-diffusible epi-metabolome (> Fig. 2a) is the only fraction of the chemical pool of the polluted site that can be the subject of the combined metabolic network of a bacterial community. The third key abstraction comes from the realization that microbial communities do contain multiple variants of enzymes that execute the same reaction on the same substrate (Witzig et al., 2006), albeit with nonidentical efficiencies (Junca and Pieper, 2004). These variants are often encoded in the same genome (duplicated or not: see, e.g., the many oxygenases borne by Rhodococcus sp. RHA1 mentioned above) as well as in different species present in the site. Most often, the first enzymes of peripheral biodegradative pathways while having an optimal specificity for a given substrate also have considerable activity on other compounds (Pazos et al., 2003). This means that the conversion of substrate A into product B in a microbial community will be the result of the additive action of all activities that can execute such a reaction, regardless of who encodes the enzyme and who benefits metabolically. The term pan-enzyme (> Fig. 3) has been proposed to designate the result of pooling all activities that bring about an identical reaction on the same substrate and originate the same product(s). The result of the three abstractions argued above is a scenario in which chemicals transit through a biodegradative landscape that integrates all possible reactions available in the site until the intermediates find its way into the central metabolism for production of biomass, or CO2 and water (> Fig. 2b). For the sake of modeling it is thus useful to visualize microbial communities behaving as a genetic/catalytic landscape that is eroded and reshaped by the flow of chemicals (> Fig. 2b). This limits the number of variables at stake to proceed with a quantitative analysis of the enzymatic potential available for cleanup of polluted sites. The University of Minnesota Biocatalysis/Biodegradation Database (UMBBD, http:// umbbd.msi.umn.edu) and its many utilities is the subject of a separate chapter of this book (> Chapter 62, Vol. 5, Part 4). The network that results from connecting all known reaction intermediates of the database in a fashion independent of the microbial host (Pazos et al., 2003) had a scale-free organization with connectivities not unlike those found in metabolic networks of single organisms. This analysis allowed the formulation of what has been called since the Global Biodegradation Network, which is the metabolic body that encompasses all known reactions that can be made by microorganisms, regardless of their origin and host species (Pazos et al., 2003). Such a network reflects the complete biodegradative potential of the global microbiota. While not all reactions might be available at any time simultaneously and some of them can even be incompatible, the concept provides a basis to the catabolic gene landscape scenario discussed above. In this Global Biodegradation Network any diffusible reaction intermediate can be shared by catabolic pathways present in different microorganisms until it finds the location in the landscape that is more proximal to the central metabolism. Such a virtual catalytic body allows the identification of all possible intermediates in the catabolism of a given chemical when passed through the merged metabolism embodied in the network. The MetaRouter system (Pazos et al., 2005) allows visualization through a web interface of all possible pathways that a large number of compounds can take through known steps of all the reactions taken from the UMBBD. This results in pathways that are a patchwork of genes/enzymes that come from different bacteria, sometimes having very

Systems Biology of Hydrocarbon Biocatalysis

. Figure 2 (Continued)

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different lifestyles. Although MetaRouter handles only compounds for which some biodegradation information is available, it provides valuable information of how given compounds could be eliminated through the merged metabolism of a complex community (Pazos et al., 2005). Apart from UMBBD and MetaRouter resources, it is possible now to make systemic analyses of the regulation of catabolic pathways on the basis of the data deposited in the system called Bionemo (http://bionemo.bioinfo.cnio.es). This platform includes information on sequence, domains, and structures for proteins related to biodegradation, including regulatory elements and transcription units for catabolic genes (Carbajosa et al., 2008). Information on regulatory proteins has been built using the results extracted from published articles and are linked to the underlying biochemical network. In its current version, Bionemo contains information on the transcriptional control of nearly 100 regulated promoters and 100 transcription factors. The system is available not only via a web server, but the full database is also downloadable as a PostgresSQL dump for other analysis tools. To the best of our knowledge, Bionemo is the most complete resource to make global analyses on transcriptional regulators from pathways that respond to hydrocarbon-related chemicals.

7

Syntrophic and Ectopic Metabolism

Biocatalysis of any given substrate S through a multistep biochemical route S ààà P (which might be CO2 + H2O) may occur through the action of a single performer microorganism, endowed with all enzymes required for complete mineralization of the compound. But, by the same token, the combined activities of various strains in a consortium may afford degradation of chemicals that cannot be performed in by single strains (de Lorenzo, 2008). The coexistence of various microbial species in the same niche, secreting part of their metabolic pool to the external medium opens two scenarios of merged metabolism. This is typical of polluted sites, in which a whole of chemical structures is often exposed to the catalytic abilities of multiple strains. One possibility is that a complex consortium includes strains that possess partial catabolic pathways for a certain substrate. Their association thus gives rise to a sort of metabolic syndicate in which one set of reactions is run by one strain (which benefits in terms of biomass buildup or energy production) and the rest are executed by one or more

. Figure 2 Abstracting key features of environmental catalysis. (a) Categories of metabolites at stake in biocatalysis. Catalytic bacteria posses a nondiffusible metabolome that – unless cells are lysed – is never secreted. Other metabolites (typically amino acids and organic acids) can diffuse out the cells but the high turnover makes their secretion very uncommon. Finally, compounds that are metabolized slowly chance to diffuse out and become available to other members of the population. The epi-metabolome is the pool of such freely diffusible intermediates. (b) The catabolic gene landscape. One can visualize and model environmental catalysis processes through a complex microbial community as the flow of epi-metabolites A!B!C!D through an uneven landscape of pan-enzymes (PE1, PE2, etc.). (c) Accessing the catabolic landscape. One experimental approach to reveal the enzymatic landscape of a given microbial community involves extraction of the meta-proteome of the place, its analysis in 2D gels, and the use of specific antibodies against different types of enzymes to pinpoint the number and abundance of each protein enzyme type.

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. Figure 3 Exposing pan-enzymes in complex environmental settings. (a) Waste treatment plant of a petroleum refinery. Some of the most interesting sites to survey microorganisms and enzymatic activities for oil biotechnology are located in the treatment facilities for residual waters of petroleum processing plants (the one shows is one of the aerobic tanks of the REPSOL factory in Puertollano, Spain. (b) Revealing 2,3-dihydroxibiphenyl dioxygenase. A small sample of the residual waters of the tanks shown above were plated on a minimal medium with citrate as the only C source, let grow and then sprayed with a 1% suspension of 2,3-dihydroxibiphenyl. Note the diversity of intensities displayed by some of the colonies with varied morphologies (surely indicating various species), which indicate the presence of 2,3-dihydroxibiphenyl dioxygenase activity. (c) One pan-enzyme is the pool of all enzymatic activities that execute the same reaction (2,3-dihydroxibiphenyl dioxygenase in this example) in a given environmental setting, regardless of the specific member of the community that carry them. The net enzymatic activity operating at the site will be the addition of each of the reactions contributed by both culturable and non-culturable members of the community.

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members of the consortium. This is the traditional syntrophic scenario (greek sym, with and trophe, nourishment) in which microorganisms of two or more different species or strains are mutually dependent on one another for nutritional requirements. Archetypical examples of this sort include the process of cellulose digestion by the microbial community of the rumen as well as the degradation of PCBs by consortia of biphenyl oxidizers and chlorobenzoate degraders. The angle of syntrophy is that partners in the process benefit metabolically from the joint process. To this end, the corresponding metabolic pathways must be regulated such that the first substrate induces the first catabolic segment in the first strain, the product of which induces a second pathway in the second strain, and so on. However, many catabolic pathways for xenobiotics are expressed constitutively or under the control of promiscuous regulators. This has been generally interpreted as the result of an ongoing and still suboptimal evolution of the transcriptional control. But a noisy regulation of this sort is likely to end up in metabolic chaos, as some pathways might be expressed in the presence of the wrong nonsubstrates and may not be produced when they are needed. If such a situation is clearly deleterious, why is it so frequently found? Does regulatory promiscuity originate new types on interactions in metabolic communities? There seem to be specific scenarios where promiscuous regulation and semi-constitutive or altogether constitutive expression or catabolic pathways could have a beneficial effect if the corresponding bacterium lives in a multi-strain community (> Fig. 4). Biodegradation of any given substrate S through a multistep biochemical route S ààà P (which can be eventually end in CO2 + H2O) can be executed by a combination of metabolic steps made present by different bacteria (1–3 in > Fig. 4). Such metabolic segments may form part of catabolic operons for substrates other than S, the result being a new pathway with contributions from enzymes located in nonrelated operons of separate bacteria and acting on the diffusible epimetabolome (see above). If the endogenous regulators/promoters of the operons that encode the enzymes were absolutely specific for each pathway and each strain, such an ectopic route (greek ek, out and topos, place) would never materialize. In contrast, if S activates promiscuously the corresponding promoters, then the community could increase its biodegradative potential. The difference between the syntrophic metabolism mentioned above and what we call here ectopic metabolism is that the benefit for the individual members of the microbial consortium would be indirect, for instance, by eliminating the toxicity of the degraded compound. In this context, it is likely that these types of environmental circumstances counteract the selective pressure toward evolving a very specific regulation for xenobioticdegrading genes. It is plausible that transcriptional factors involved in the process and the regulatory networks to which they belong are plastic enough to move back and forth between various degrees of promiscuity/specificity, constitutive/inducible expression, and more or less physiological coupling, depending on environmental conditions and the type of partners in the consortium. These scenarios have fundamental and practical consequences that deserve further studies with the tools of Systems Biology.

8

Predicting Catalysis and Biodegradation

Given the large number of compounds of environmental interests, different approaches have been entertained since the late 1990s to set up systems that can predict the fate of chemical compounds before testing experimentally the capacity of the microbiota to degrade them. One useful procedure focuses on the reactivity of functional groups present in the molecules to be

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. Figure 4 Ectopic metabolism caused by promiscuous regulation. The scenario represented in the figure involves three strains (1, 2, and 3), which do not degrade separately compound S because it is not a growth substrate of their separate catabolic abilities encoded in strain-specific operons. However, strain 1 contains an operon that includes one enzyme (A) that can take S into an intermediate product , which can be transformed into by the enzyme B of strain 2, which in turn can be further metabolized or altogether destroyed by enzyme C of strain 3. If the respective regulators of each operon (R1, R2, and R3) are completely faithful to the distinct growth substrates of each of the pathways, then S cannot be degraded by the consortium. However, if S induces gratuitously the operons owing to the promiscuity of R1, R2, and R3, the compound can become ultimately degraded despite not being the growth substrate of any of the strains.

degraded (Ellis et al., 2008; Fenner et al., 2008; Hou et al., 2003). The properties and reactivities of such groups are then implemented into a number of rules assigned to individual chemical reactions that provide a stepwise prediction on whether given discrete reactions that involve functional groups are likely to occur on the molecule under scrutiny. The system allows visualization of the chemical structures of all the potential reaction intermediates for each virtual pathway. Unfortunately, the straight reactivity of functional groups in a molecule may not be the ultimate determinant of the environmental fate of the compounds under scrutiny. As shown in > Fig. 5, biodegradation (more so in actual bioremediation scenarios) implies one or more pre-catalysis steps as well as post-catalysis, downstream consequences of metabolizing any given compound. The only functioning of the actual catalytic step(s) of a biodegradative pathway does not guarantee removal of the target compound. Peripheral metabolic pathways which are typical of biodegradation routes need to be satisfactorily coupled to the central metabolism and to the overall energy balance of the cells. Ideally, biodegradation should be linked to growth or detoxification in order to provide a selective advantage to the cells that bear the catalytic activity (de la Pena Mattozzi et al., 2006). But, unlike the chemical and biochemical aspects, such physiological facets of biodegradation are more difficult to implement in a predictive system. An illustrative example in this sense is the inability of strain B. xenovorans LB400 to degrade completely PCBs despite having in its genome all genes, which in principle are necessary to this end (Perez-Pantoja et al., 2008). Typically, B. xenovorans LB400 converts PCBs into chlorobenzoates and stops there. But the bacterium has at least one system for

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. Figure 5 Biotic and abiotic steps in microbial biocatalysis. One key difference between chemical catalysis and its live (micro)biological counterparts is the number of factors at play in the process. There is first the issue of bioavailability, as the substrate of the desired reaction (A) may not access directly the core catalyst (often an intracellular enzyme) because of its intrinsic lack of solubility or its adsorption to mineral matrixes. In addition, the substrate might also be a stressor for the live catalysts. Second, the desired transformation step may or may not be coupled to growth and may originate side products (C, F) with deleterious effects on the microorganism. Finally, the product of the process (Z) might be toxic and inhibit the progression of the whole biotransformation. This scenario provides an optimal chance to apply a Systems Biology approach to the design of processes of this sort.

catabolism of clorobenzoates, which is perfectly active when such compounds are given as growth substrates (Martinez et al., 2007). Biochemically, the strain can thus mineralize PCBs completely, but a number of physiological problems prevent it to do so in vivo (Agullo et al., 2007). Along the line, although the early enzymes encoded by the bph system of this strain have activity on a large variety of PCBs, B. xenovorans LB400 can grow only on plain biphenyl and lightly chlorinated PCBs (Pieper and Seeger, 2008). One possibility to overcome this problem is to set aside chemical reactivity and simply train a rule-based classification system for detecting the association between certain chemical compound descriptors and environmental fates (Gomez et al., 2007). Such descriptors are based on the deconstruction of chemical structures in atomic triads (also referred to as chemotopes) plus two additional qualities (molecular size and solubility). A machine learning system was then used to identify explicit rules that associate compound vectors to environmental fates as inferred from the analysis of the Global Biodegradation Network discussed above. The examination of many compounds with this system suggests that the frequency of atomic triad sets the fate of the molecules when facing the Global Biodegradation Network. Whether based on known chemical reactivity, on machine learning approaches or on both, biodegradation prediction systems will be invaluable to provide criteria for guiding bioremediation interventions.

9

Outlook: From Systems to Synthetic Biology

The growing availability of genomic sequences of microorganisms that have activity on petroleum compounds and their derivatives provides a start to the complete comprehension of their interactions. But having a catalog of genes, proteins, and metabolites is not enough and conceptual frames imported from Physics, Engineering, and Information Theory are required to move a descriptive science into a more quantitative and predictive discipline.

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Systems Biology exactly fills that interface between Biological information, often lacking any format, and quantitative knowledge. As discussed above, complexity is not an absolute quality of systems, but a mere comparative judgment of the number of elements and connections that participate in any given relational body. In that respect, hydrocarbon biotechnology could be visualized as a complexity pyramid that goes from singular genes and operons encoding catalytic functions of interest, all the way to a complex and highly dynamic microbial community operating on an extensive landscape (de Lorenzo, 2008). Depending on the specific agendas, the various levels of complexity have to be addressed with diverse conceptual frames and technologies. While programming biocatalysis in a reactor with a defined medium asks for the handling of a set of genes in a specific strain, the planning of environmental interventions for removal of toxic waste or accidental spills, and shifts the focus into the chemical, abiotic, and catabolic landscape of the place. In this respect, Systems Biology provides tools to cover all the scales of complexity that are needed to make processes of this sort more reliable. But how? One way to modify the behavior of a system is to rewire artificially a subset of interactions within the relational body. As mentioned above, from the mid-1980s up to the late 1990s numerous attempts were made to alter such interactions by designing genetically modified microorganisms for environmental release as agents for bioremediation and control of organic pollutants. Yet, the field eventually came to a standstill after recurrent failures to program bacteria to behave in a nonnatural fashion in scenarios quite different of the controlled conditions of the Laboratory. Part of the problem can be traced to the naivety of the genetic engineers of the time, who often overlooked design principles for complex systems and circuits that are commonplace 20 years later. This situation is rapidly changing owing to the expansion of Synthetic Biology and its emphasis in robust design concepts, e.g., modularity, orthogonality (i.e., context-independency, see above) and definition of systems boundaries (Arkin, 2008; Canton et al., 2008). Although the expression Synthetic Biology (SB) has been present in the scientific and technical literature since 1912 (Leduc, 1912) only in more recent times it has shifted to being an umbrella concept at the interface between Molecular Biology and sensu stricto Engineering. SB is becoming an increasingly inclusive concept, which (1) encompasses new theoretical frameworks that approach biological systems with the conceptual tools and the descriptive language of Engineering, (2) addresses standing questions with fresh approaches inspired in electric circuitry and mechanical manufacturing, and (3) pursues the creation of new materials with a´ la carte properties based on the rational combination of standardized biological parts decoupled from their natural context. In fact, standardization and detailed description of minimal biological parts and their interfaces, to the degree of reliability of the components of modern electronic circuits is one of the trademark of the whole field. The basic notion behind SB is that any biological system can be seen as a combination of functional, stand-alone elements not unlike those found in man-made devices, and can thus be disclosed as a limited number of components. These can then be reconstructed in novel configurations for the sake of modifying existing properties or creating altogether new ones. Biological properties and activities can be – under the conceptual frame of SB – abstracted, made modular and rewired with a defined connectivity and precise systems boundaries (de Lorenzo and Danchin, 2008). Although the SB agenda is still in an early stage at the time of writing this article, the field is likely to change traditional practices in biocatalysis – both in the reactor and in the field. De novo synthesis of complete bacterial genomes is becoming technically feasible and the possibility to design microbial catalysts a´ la carte is now much closer.

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Acknowledgments The work made in Authors’ Laboratory was supported by research grants of the Spanish Ministry of Science and Innovation, by contracts of the Framework Programme of the EU and by Funds of the Autonomous Community of Madrid.

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Cellular Ecophysiology: Problems of Hydrophobicity, Bioavailability

42 Introduction: Problems of Hydrophobicity/ Bioavailability H. Harms1,* . K. E. C. Smith2 . L. Y. Wick1 Helmholtz-Zentrum fu¨r Umweltforschung, Department of Environmental Microbiology, Leipzig, Germany *[email protected] 2 Department of Environmental Chemistry and Microbiology, The National Environmental Research Institute, Roskilde, Denmark

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1 Introduction: The Hydrophobic Effect and its Implications for Hydrocarbon Microbiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1440 2 The Bioavailability of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1441 3 Living with Poorly Bioavailable Substrates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1444 4 The Bioaccessibility of Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1445 5 Influences of Chemical Hydrophobicity on Bioavailability and Bioaccessibility . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1446 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1449

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_98, # Springer-Verlag Berlin Heidelberg, 2010

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Introduction: Problems of Hydrophobicity/Bioavailability

Abstract: This chapter attempts to show how the hydrophobicity of hydrocarbons influences their availability for toxic exposure, microbial degradation and growth, and how it can control the carrying capacity of a microbial habitat. Bioavailability is operationalized and presented as a process at the interface between biological dynamics and physicochemical constraints.

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Introduction: The Hydrophobic Effect and its Implications for Hydrocarbon Microbiology

Hydrocarbons have in common that they are repelled or expulsed from a mass of water. This characteristic of chemicals is referred to as hydrophobicity (from the Greek hydro- for water and phobos for fear). Hydrophobicity has its physical origin in the polar nature of water and the tendency of water molecules to form hydrogen bonds with each other. Hydrogen bond formation between water molecules is energetically more favourable than the interaction of water molecules with non-polar (non-hydrogen bond forming) molecules or phases (Schwarzenbach et al., 2005). Water thus repels or expulses hydrophobic chemicals in favor of bonding with itself. From a thermodynamical point of view this can be ascribed to the high entropic cost of forming a cavity inside a water mass around nonpolar molecules. The term hydrophobicity is thus misleading as the force giving rise to the so-called hydrophobic effect arises from the hydrophilic partner in the interaction. Hydrophobicity most frequently, but not always, goes along with lipophilicity, i.e., the tendency of a molecule to partition into lipids or other non-polar organic phases and to accumulate therein. Hydrophobicity as such is difficult to measure and to describe in quantitative terms. A convenient proxy for a compound’s hydrophobicity is its tendency to partition between water and a liquid organic phase. To allow comparison, octanol has been chosen as the conventional organic solvent used to quantify the hydrophobicity of many kinds of chemicals. The octanolwater partition coefficient KOW is the ratio of the concentration of a compound in an octanol phase to its concentration in an adjacent water phase at equilibrium. The KOW is typically determined in partition experiments, but there are now methods for the calculation of KOW from the chemical structure available as free internet resources. A practical example is the Sparc On-line Calculator (http://ibmlc2.chem.uga.edu/sparc). Observed KOW values span such a wide range that conventionally Log KOW values are reported. Chemicals possessing values of KOW higher than 1 can be regarded as hydrophobic because they prefer the organic phase to the aqueous phase. Hydrocarbons generally have Log KOW above 3. In > Fig. 1 octanol-water partition coefficients of a series of environmentally relevant hydrocarbons are presented together with two other observable expressions of hydrophobicity namely the tendency to become volatilized (become expulsed from water into an air phase) and low water solubility (pure substance being repelled by water). From > Fig. 1, it becomes obvious there are some general rules for the degree of the hydrophobicity of hydrocarbons as a function of their structures. The hydrophobicity of hydrocarbons rises with the molecular mass and the degree of saturation. Long alkane chains are thus more hydrophobic than short ones, saturated aliphatics more hydrophobic than unsaturated aliphatics and aromatics of the same molecular mass, and polycyclic aromatic hydrocarbons consisting of a higher number of rings are more hydrophobic than low molecular mass PAHs. In the environment hydrocarbons can be present as individual molecules in the gaseous, water-dissolved, surface-adsorbed or matrix-absorbed form or as separate bulk phase (liquid or solid) that either floats on water, forms blobs or aggregates in pores of sediments and soil or spreads as thin layers on biota.

Introduction: Problems of Hydrophobicity/Bioavailability

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. Figure 1 Plots of water solubility (Cwsat), octanol-water partition coefficient (KOW), and dimensionless Henry coefficient (H’) versus molecular mass for four typical groups of hydrocarbons: PAHs, methylated PAHs, n-alkanes and branched alkanes. Typical low, middle and high mass compound from the PAHs and n-alkanes have been marked on the plots using numbers (PAHs: 1 naphthalene; 2 phenanthrene; 3 benzo(ghi)perylene. n-Alkanes: 4 n-pentane; 5 n-decane; 6 hexacosane). Data from Schwarzenbach et al., 2005; Eascott et al., 1988, and http://www.lec. lancs.ac.uk/ccm/research/database/index.htm.

2

The Bioavailability of Hydrocarbons

Whenever microorganisms interact with hydrocarbons, the hydrophobic effect plays a role. Active microbes are typically surrounded by water and their interaction with hydrocarbons will involve the water phase. The hydrophobicity of a chemical is thus often taken as an

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indicator for its bioavailability, in particular for microorganisms, which are believed to take up chemicals as water-dissolved molecules. The term bioavailability is generally used to refer to the degree of interaction of chemicals with living organisms. Unfortunately none of the existing further refinements of this all-encompassing definition are generally accepted and it appears that a short definition of practical value cannot be given (NRC, 2003; Harmsen, 2007), but bioavailability needs to be defined starting from a generally accepted concept to its specific use (ISO 17402). It appears thus useful to start out from this small common concept and ground and develop an operational definition of bioavailability in the context of hydrocarbon microbiology before beginning to scrutinize the various bioavailability processes that govern the interactions of microbes with hydrocarbons. Obviously, quantification requires that bioavailability has a physical dimension. As there is no general agreement on a dimension or unit of bioavailability either, we need to ask what an operational dimension of bioavailability could be? Before we answer this question, it is important to distinguish between bioavailability for microbial degradation on the one hand and for bioaccumulation on the other hand (> Fig. 2). (1) Microbial hydrocarbon degradation is a consumptive process that tends to strive towards an at least transient steady state of biological hydrocarbon uptake and environmental hydrocarbon re-provision (cell 1 in > Fig. 2). It has been shown that the rate of substrate diffusion to an organism and its rate of uptake by the organism tend to become equal (Koch, 1990).

. Figure 2 Illustration of the difference between chemical bioavailability to a cell actively degrading a dissolved hydrocarbon (Cell 1) and to a cell passively accumulating hydrocarbons (Cell 2). Note that cell 1 may behave with respect to other hydrocarbons in a mixture like cell 2. The water-dissolved cell surface exposure concentrations Cw cs, the hydrocarbon transfer flux (only present in cell 1 as indicated by the arrow) and the hydrocarbon concentration in the cells (visualized as different shades of grey) differ substantially. Cw and Cw eq are the water-dissolved concentration and the water-dissolved concentration at equilibrium, respectively.

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The hydrocarbon concentration to which the microbe is exposed and the actual rate at which the hydrocarbon enters the organism are controlled by biological kinetics and environmental mass transfer. The steady-state mass transfer rate is determined by the specific affinity (i.e., the slope of the first order part of the whole-cell Michaelis–Menten curve) of the cell for the chemical in question and the water-dissolved concentration in contact with the cell. Microorganisms having a high specific affinity for their substrate can reduce the exposure concentration to extremely low values, because they take up their substrate so rapidly (drive such a fast substrate flux) that they create a steep concentration gradient between their surface and the bulk water at some distance. (2) The situation is different for a microbe that is subject to the toxic effects of a bioaccumulating compound without having the possibility to degrade it (cell 2 in > Fig. 2). Here, an equilibrium situation will be approached. The catabolically inactive microorganism will be subject to passive inflow and bioaccumulation of the chemical. With time, equilibrium will be reached that is characterized by firstly, a hydrocarbon content of the organism that is mainly controlled by the microbe’s lipid content and the lipophilicity of the hydrocarbon, secondly, exposure of the cell surface to the aqueous equilibrium concentration, and thirdly, the absence of a net flux of hydrocarbon into the organism. Unlike the steady state observed for biodegradation (characterized by a low exposure concentration and high mass transfer), the endpoint of bioaccumulation is thus characterized by a relatively high aqueous equilibrium concentration and zero mass transfer. Bioavailability for passive bioaccumulation can therefore be quantified by approaches addressing the chemical activity of a compound, such as equilibrium extraction using solid phase microextraction (SPME) fibers (Reichenberg and Mayer, 2006). In the following we will see that it is more difficult to operationalize biovailability for biodegradation. This is due to the dynamics of biodegradation and the frequently long duration over which biodegradation needs to be sustained by bioavailable compound. From a practical perspective (e.g., in bioremediation or industrial biotransformation) we are interested in reaction rates and reaction endpoints, e.g., because remediation goals need to be matched in reasonable time scales. The reaction rate appears thus to be an immediate measure and appropriate dimension of bioavailability. However, there are some problems with this view since the measured rate may be limited by the capacity of the organisms to degrade or transform the chemical. One might say that the bioavailability is even higher than is apparent from the degradation rate since the mass transfer capacity remains partly unexploited. Bosma et al. (1997) have therefore defined bioavailability as the ratio of the rate at which a volumetric unit of the environment can theoretically provide a chemical to organisms to the rate at which the microbes present in this volume can theoretically degrade the compound. As these rates (or capacities) of consumption and re-provision have equal units, a dimensionless bioavailability Ba number was proposed. Values of Ba above 1 indicate degradation rates predominantly controlled by the degradation capacity of the organisms (i.e., high bioavailability), whereas values of Ba below 1 indicate degradation rates predominantly controlled by the mass transfer capacity of the environment (i.e., low bioavailability). Recently, Thullner et al. (2008) have simplified the above concept by defining bioavailability as the ratio of the actual biodegradation rate of an extant microbial community to its degradation capacity. Both concepts account for the fact that exposure concentrations are inappropriate descriptors of bioavailability since they can be very low even at high degradation rates (cell 1 in > Fig. 2). Bioavailability according to both definitions depends on how the habitat can compensate for substrate degradation. This capacity of the habitat will depend on the physical state of the chemical (dissolved, sorbed, separate phase, occurrence as individual substance vs. mixture,

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etc.), its physical characteristics (hydrophobicity, effective diffusivity, etc.) and its spatial distribution relative to the catabolically active biota.

3

Living with Poorly Bioavailable Substrates

Biodegradation requires that catabolically active microorganisms grow (or grew). The relationship between biodegradation and growth is however more complicated than often thought. In particular, the Michaelis-Menten relationship describing degradation rates and the Monod relationship describing growth are often mixed up or taken as equivalent. This is especially tempting when the former is applied to the kinetics of entire cells. It appears thus necessary to explain some important differences before using extended Monod kinetics to explain the constraints of life with poorly bioavailable substrates. Both relationships are presented in the form of hyperbolic plots of activity or growth against the substrate concentration. The equations of the general form a = b * c/(d + c) are mathematically equivalent, where a is the actual rate, b the maximum rate (the biological capacity), c the concentration and d the concentration resulting in half maximal rate. The similarity hides that the MichaelisMenten equation is a mathematically derived relationship whereas the Monod relationship is an empirical model that has been found to fit growth data. It also hides that the experimental verification of Michaelis–Menten is performed with constant biological materials (pure enzyme, a raw protein extract or resting cells), whereas Monod data are obtained in chemostat cultures that are slowly shifted from one steady state substrate concentration to the next one, shifts that allow adaptation and regulation events to occur (Kovarova-Kovar and Egli, 1998). Even more important for our discussion of bioavailability is another difference between both concepts. Whereas Michaelis–Menten enzyme kinetics apply down to minute concentrations, since even the last substrate molecule has the chance to meet an enzyme (or a transporter on a cell surface), the classical Monod concept disregards the fact that minute substrate concentrations corresponding to minute substrate fluxes do not provide enough energy to organisms to allow them to grow. A certain critical substrate flux will correspond to an exposure concentration that will only be sufficient for the maintenance of the existing cells and even lower concentrations will even lead to the die-off of existing cells. This offset of the hyperbolic relationship has been accounted for by a maintenance rate coefficient extending the classical Monod-kinetics (Van Uden, 1967), which is equivalent to the rate of culture decay (also termed negative growth) at substrate concentration zero (> Fig. 3). A consequence of this offset is that in many environments, populations will grow as long as the substrate flux allows them to do so, thereby approaching a situation where the substrate flux only allows for maintenance of the extant population. This has been shown for cultures in chemostats with biomass retention (Tros et al., 1996), in biofilms growing on poorly soluble anthracene crystals (Wick et al., 2001) and for bacteria relying on the diffusion of naphthalene from spatially separate sources (Harms, 1996). Under conditions of reduced bioavailability (i.e., high bias between the cell exposure concentration and the total concentration in the system), even high total concentrations may allow for the maintenance of only relatively small populations, which in turn bring about only slow and sometimes non-detectable degradation progress (> Fig. 4). In ecological terms, substrate bioavailability controls the carrying capacity of microbial habitats, i.e., their capacity to maintain a population. The maintenance requirements have consequently been

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. Figure 3 Monod-plot accounting for the maintenance energy requirements of bacteria (a) and visualization of different substrate consumption-dependent concentration gradients around bacteria in habitats permitting different rates of substrate transfer (b). Low substrate transfer in organic-rich soil (bottom graph in B) reduces the cell exposure concentration (C2 in A and B) to below the threshold for population maintenance, whereas high substrate transfer in sand (top graph in B) sustains a cell exposure concentration (C1 in A and B) permitting population growth.

proposed as one reason for the observation of remediation thresholds particularly in environments that permit only slow substrate mass transfer (Bosma et al., 1997).

4

The Bioaccessibility of Hydrocarbons

The previous discussion focused largely on the degradation rate. However, reaction endpoints can not been predicted from bioavailability that is defined in a way that it is linked to actual, ephemeral degradation rates. The capacity of an environment to provide a chemical at a constant rate may become exhausted because certain labile pools of the chemical may become emptied. This possibility has inspired the differentiation of an immediate bioavailability (compound at hand actually being taken up) from a potential bioavailability named bioaccessibility (compound that may become bioavailable by dissolution, desorption, diffusion etc.) (Semple et al., 2004). In this concept, the bioaccessible compound has the dimension of a fraction of the total compound and is thus an appropriate descriptor of the possible degradation endpoint. The bioaccessible fraction comprises the chemical mass that can reach the biota (or vice versa) within a predefined time frame. The distinction between bioavailability and bioaccessible is to some extent equivalent to that between a compound’s chemical activity as a quantitative descriptor of spontaneously mobilizable compound and an accessible quantity

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. Figure 4 Scheme explaining the relationship between the mass transfer capacity of a microbial habitat, the carrying capacity of this habitat and the observed residual concentration of a hydrophobic chemical. Low substrate transfer in organic-rich soil (upward arrow of case 1) limits the carrying capacity of this habitat to low cell numbers (upper arrow pointing to the right), which are insufficient for observable contaminant degradation although residual concentrations are still relatively high (upper arrow pointing to the left). High mass transfer in sand (case 2) increases the carrying capacity thereby facilitating the reduction of residual chemical.

that has to be operationally defined from case to case as introduced by Reichenberg and Mayer (2006). These authors propose equilibrium sampling devices for the quantification of the chemical activity aspect of bioavailability and mild extraction methods for the quantification of the fraction of bioaccessible compound.

5

Influences of Chemical Hydrophobicity on Bioavailability and Bioaccessibility

Various phenomena of relevance for microbe-hydrocarbon interactions that arise from a compound’s hydrophobicity are presented in > Table 1. Effects on the molecular level such as low water solubility and the tendency of hydrophobic compounds to sorb to surfaces or partition into animate or inanimate materials can be distinguished from bulk phase-related phenomena such as immiscibility of phases and low wettability of hydrophobic phases. Only short descriptions will be given here since many of these phenomena will be treated in more detail in subsequent chapters. The low water solubility controls the maximum substrate concentration or energy density in the aqueous environment of the microbes. Unless microorganisms take up hydrocarbons

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. Table 1 Environmentally relevant phenomena associated with the hydrophobicity of hydrocarbons

Phenomenon

Associated phenomena and environmental expression

Low water solubility

Relevance for microbe-hydrocarbon interaction Low substrate/energy density of the aqueous habitat Low toxicity

Sorption

Adsorption to solid surfaces

Reduced bioavailability in the bulk water Lowered toxicity High substrate density at the solid surface

Partitioning

Pore condensation

Disappearance in pores inaccessible for microbes

Accumulation in air-water interfaces

Reduced bioavailability in the bulk water

Absorption in particulate and dissolved organic matter

Reduced bioavailability due to lower aqueous concentration

High substrate density at the air-water interface

Increased bioaccessibility when the organic matter is mobile Bioaccumulation

Increased substrate density in organisms Increased toxicity

Dissolution in micelles and NAPLs Formation of Minimized interfacial area immiscible separate phases Floating of light non-aqueous phase liquids (lNAPLs)

Reduced bioavailability in the bulk water Increased bioaccessibility due to micelle mobility Small area for settlement and microbial attack Slow dissolution limiting the re-supply of consumed substrate in the aqueous phase Shielding of oxygen Lowered bioaccessibility due to heterogeneous substrate distribution Limited provision of e-acceptors and nutrients

Sinking of dense non-aqueous Heterogeneity of substrate distribution phase liquids (dNAPLs) Limited provision of e-acceptors and nutrients

Low wettability

Low bulk mobility due to viscosity and sequestration in pores

Low bioaccessibility

Instability of emulsions

Reduced area for settlement and microbial attack

Beading of water on HC phase Reduced contact of HC with the aqueous in unsaturated systems habitat of microbes

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directly from a non-aqueous phase (a possibility that will be discussed in > Chapter 3), or cosolvents increase the water solubility (Wick et al., 2002), the water solubility will cut off a population’s Michaelis-Menten curve thus setting an upper limit to the specific degradation rate. On the positive side low solubility reduces a chemical’s toxicity to microorganisms. Sorption to solid surfaces and accumulation in the air-water interface (Hoff et al., 1993) will reduce the aqueous concentration with the consequences discussed before. One can distinguish a first case where the sorbing interfaces are exposed to bacteria and a second case where the sorbent is present in pores too small to be accessed by bacteria. Increased substrate and energy densities arise at these interfaces. Transfer from the bioaccessible to the bioavailable state, however, will require desorption. At equilibrium the chemical activities of the sorbed compound and its dissolved counterpart will be the same, meaning that microorganisms will not benefit from higher concentrations at interfaces (because they represent bioaccessible and bioavailable compound) but only from the short transport distance of desorbing chemicals (Van Loosdrecht et al., 1990). Except for the second case that molecules need to diffuse out of pores, the overall influence of adsorption on bioavailability will be relatively low, due to the immediacy of the release steps. Partitioning into inanimate phases, also referred to as absorption, has more drastic consequences for bioavailability, since it spatially separates the chemical from the aqueous phase. Molecules entering organic matter (OM) will travel towards the centre of the OM until partition equilibrium is achieved. To release them again may take a long time, since the same distance will have to be passed again on the way out, notwithstanding that chemical changes can occur that strongly retain the molecules (Luthy et al., 1997). Partitioning into small entities of OM such as the so-called dissolved organic carbon can have the effect that the chemical is absorbed, but in a highly mobile form of OM that may act as a vector and facilitate its transfer to biota. The same holds true for surfactant micelles. Partitioning into the microbes themselves obviously concentrates the chemical at the point where it can be consumed or act as a toxicant. The formation of separate hydrocarbon phases that are immiscible with water has consequences both on the microscopic and on the macroscopic scales. On the microscopic scale it reduces the interfacial area through which hydrocarbon molecules are released into the water phase. This limits the dissolution rate thereby affecting bioavailability via the substrate reprovision rate. The minimized interface also limits the area available for settlement of microorganisms. As only few bacteria will be located in the proximity of the hydrocarbon phase, immiscibility increases the mean distance between the hydrocarbon mass and most of the biota in the system. One can also say that it increases the heterogeneity of the hydrocarbon distribution. Measures such as emulsification help to overcome this effect. In soils and sediments the formation of viscous non-aqueous phases also has consequences for the bulk movement of hydrocarbons, which tend to stick inside of pores, a phenomenon known from the difficulty to extract residual oil from oil reservoirs. On the macroscopic scale non-aqueous phase liquids, depending on their gravimetric density may form layers on top of the groundwater surface (or the ocean) or sink down to the confinement layers (or ocean sediment). In both cases the ratio of the NAPL-water interfacial area to the NAPL mass is very low and considerably restricts the transfer of bioaccessible into the bioavailable form. In the case of light NAPLs it may in addition shield oxygen or nutrients form the biota sitting below the NAPL thereby affecting the bioavailability of these important factors. Finally, the low wettability plays a role where an air phase is present that competes (and succeeds in the competition) for the formation of an interface with hydrocarbons. Examples

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can be found in the vadose zone of terrestrial environments as well as the beading of raindrops on the waxy surfaces of leaves.

6

Research Needs

Future research into hydrocarbon bioavailability should lead to the development of reliable methods for the prediction of bioaccessibility (ideally a small set of tests tailored for a corresponding set of pollution situations). This will require an improved understanding of physico-chemical bioavailability processes, particularly those prevailing in complex hydrocarbon mixtures and the elucidation of microbial strategies to improve bioavailability under these conditions. The latter should also address possible roles of microbial community members not directly involved in hydrocarbon degradation. An example is the function of fungal mycelia as facilitators of bacterial movement treated in Chapter 10. Another largely neglected aspect is the capability of hydrocarbon degrading populations and communities to decouple their biomass build-up and survival from the hydrocarbon provision by either cometabolism or oligotrophic multi-substrate utilisation. Finally, research into energy-efficient in situ methods for bioavailability enhancement should be conducted.

References Bosma TNP, Middeldorp PJM, Schraa G, Zehnder AJB (1997) Mass transfer limitation of biotransformation: Quantifying bioavailability. Environ Sci Technol 31: 248–252. Eascott L, Shiu YS, Mackay D (1988) Environmentally relevant physical-chemical properties of hydrocarbons: A review of data and development of simple correlations. Oil Chem Poll 4: 191–216. Harms H (1996) Bacterial growth on distant naphthalene diffusing through water, air, water-saturated and nonsaturated porous media. Appl Environ Microbiol 62: 2286–2293. Harmsen J (2007) Measuring bioavailability: from a scientific approach to standard methods. J Environ Quality 36: 1420–1428. Hoff JT, Mackay D, Gillham R, Shiu WY (1993) Partitioning of organic-chemicals at the air water interface in environmental systems. Environ Sci Technol 27: 2174–2180. ISO (2008) ISO 17402 – Soil quality – requirements and guidance for the selection and application of methods for the assessment of bioavailability of contaminants in soil and soil materials. Koch AL (1990) Diffusion – the crucial process in many aspects of the biology of bacteria. Adv Microb Ecol 11: 37–70. Kovarova-Kovar K, Egli T (1998) Growth kinetics of suspended microbial cells: From single-substratecontrolled growth to mixed-substrate kinetics. Microbiol Mol Biol Rev 62: 646–666.

Luthy RG, Aiken GR, Brusseau ML, Cunningham SD, Gschwend PM, Pignatello JJ, Reinhard M, Traina SJ, Weber jr. WJ, Westall JC (1997) Sequestration of hydrophobic organic contaminants by geosorbents. Environ Sci Technol 31: 3341–3347. Committee NRC (2003) NRC Committee on Bioavailability of Contaminants in Soils and Sediments. Bioavailability of contaminants in soils and sediments: processes, tools and applications. Washington DC: The National Academic Press. Reichenberg F, Mayer P (2006) Two complementary sides of bioavailability: Accessibility and chemical activity of organic contaminants in sediments and soils. Environ Toxicol Chem 25: 1239–1245. Schwarzenbach RP, Gschwend PM, Imboden DM (2005) Environmental Organic Chemistry. New York: Wiley. Semple KT, Doick KJ, Jones KC, Burauel P, Craven A, Harms H (2004) Defining bioavailability and bioaccessibility of contaminated soil and sediment is complicated. Environ Sci Technol 38: 228A–231A. Thullner M, Kampara M, Harms H, Wick LY (2008) Impact of bioavailability restrictions on microbially induced stable isotope fractionation: 1. Theoretical calculation. Environ Sci Technol 42: 6544–6551. Tros ME, Bosma TNP, Schraa G, Zehnder AJB (1996) Measurement of minimum substrate concentration (S-min) in a recycling fermenter and its prediction from the kinetic parameters of Pseudomonas sp. strain B13 from batch and chemostat cultures. Appl Environ Microbiol 62: 3655–3661.

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Van Loosdrecht MCM, Lyklema J, Norde W, Zehnder AJB (1990) Influences of interfaces on microbial activity. Microbiol Rev 54: 75–87. Van Uden N (1967) Transport-limited fermentation in the chemostat and its competitive inhibition: A theoretical treatment. Archiv Mikrobiol 58: 145–154.

Wick LY, de Munain AR, Springael D, Harms H (2002) Responses of Mycobacterium sp. LB501T to the low bioavailability of solid anthracene. Appl Biotechnol Microbiol 58: 378–385. Wick LY, Colangelo T, Harms H (2001) Kinetics of masstransfer-limited growth on solid PAHs. Environ Sci Technol 35: 354–361.

43 Water-Hydrophobic Compound Interactions with the Microbial Cell E. M. McCammick1 . V. S. Gomase2 . T. J. McGenity3 . D. J. Timson1 . J. E. Hallsworth1,* 1 School of Biological Sciences, MBC, Queen’s University Belfast, Belfast, Northern Ireland *[email protected] 2 Department of Bioinformatics, Padmashree Dr. D. Y. Patil University, CBD Belapur, Navi Mumbai, India 3 Department of Biological Sciences, University of Essex, Colchester, UK 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1452 2 Water as a Chaperone . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1453 3 Cytosol-Lipid Bilayer-Extracellular Solution; an Aqueous Continuum . . . . . . . . . . . . 1454 4 Behavior of Hydrophobic Substances in the Cellular System . . . . . . . . . . . . . . . . . . . . 1456 5 Do Hydrocarbons Induce Asymmetrical or Symmetrical Stresses? . . . . . . . . . . . . . . . . . 1459 6 Microbial Cells in Hydrophobic Environments: Situational-Functional Considerations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1461 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1462

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_99, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: The structural interactions of biological macromolecules, their biochemical activities and, ultimately, the metabolic function of cellular systems are dependent upon weak inter- and intra-molecular forces such as hydrogen bonds, Van der Waals forces, and the hydrophobic effect. Water molecules, and those of hydrophobic substances such as hydrocarbons, can take part in and/or modify these interactions and thereby determine the operational and structural stability of the microbial cell and its macromolecular systems. We explain how the cytosol, plasma membrane and the extracellular solution form a material and energetic continuum; and discuss the behavior of hydrophobic substances of extracellular origin as they migrate into the plasma membrane and into the cell’s interior. The adverse effects of substances with a log Poctanol-water 2, that partition into the hydrophobic domains of biological macromolecules, are discussed in relation to microbial cell function; and we speculate whether the cellular stress that they induce is symmetrical or asymmetrical in nature. In the context of the microbial environment, we take a situational-functional approach to consider how hydrophobic stressors interact with the microbial cell, and what types of evasion tactics microbes can employ to minimize their inhibitory activities. Finally, we discuss the ecological implications of hydrocarbon-induced cellular stress for microbial systems.

1

Introduction

The microbial cell is a fragile aqueous system, and dissolved substances represent potentially lethal challenges to its macromolecular structures. Microbes in Nature are exposed to an indefinite number of extracellular compounds (see > Vol. 1, Part 3), some of which will penetrate the cellular membrane. These may be utilized for metabolic purposes (e.g., for signaling; as nutrients used during structural growth or to synthesize compatible solutes), may act as toxicants that inhibit metabolism via a site-specific mode-of-action (see > Vol. 2, Part 8), and/or may induce cellular stress via physicochemical interventions in macromolecular structures and their interactions. The ecology of a microbial species, as well as its evolutionary trajectory, are determined by its ability to survive, respond, and adapt to environmental challenges, the most powerful of which are those that induce diverse forms of water stress (Brown, 1990; Hallsworth et al., 2007; Kashangura et al., 2006). All dissolved substances, even stress protectants, can act as cellular stressors that reduce water activity, induce osmotic or matric stress, impair the functionality of lipid bilayers and/or exert chaotropic activity towards biological macromolecules (Brown, 1990; Duda et al., 2004; Ferro Fonta´n and Chirife, 1981; Hallsworth and Magan, 1994; Hallsworth, 1998; Hallsworth et al., 2003, 2007). The extent to which extracellular solutes impact on microbial-cell infrastructure and metabolism is a function of the way in which their physical and chemical properties determine behavior upon/after entry into the plasma membrane. Polar solutes that hydrogen-bond with water more strongly than water molecules hydrogen-bond with themselves (i.e., kosmotropes, see Washabaugh and Collins, 1986) characteristically have a substantial hydration shell and large effective molecular radius. Such species, for example Na+ and sucrose, are generally excluded from the aqueous phases of lipid bilayers and most other macromolecule structures (Arakawa and Timasheff, 1985) and therefore induce osmotic stress (see Brown, 1990). By contrast polar compounds such as ethanol, urea and phenol that hydrogen-bond more weakly with water than water molecules do with themselves can typically move into the aqueous phase of proteins, membranes and other cellular structures. Such compounds weaken the non-covalent interactions of cellular macromolecules and thereby disorder macromolecular structures, and assemblages (i.e., they are chaotropic), inducing a non-osmotic form of water stress

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(Hallsworth, 1998; Hallsworth et al., 2003). Hydrocarbons that are sufficiently hydrophobic to preferentially partition into the hydrophobic interior of membranes or other macromolecular systems typically have a log Poctanol-water 2 (e.g., benzene and hexane; Bhaganna and Hallsworth, unpublished data). The severity of the adverse effects of hydrophobic hydrocarbons on macromolecular and cellular function is determined, in part at least, by what has been variously referred to as their hydrophobicity, lipophilicity or solventgenicity (e.g., Sikkema et al., 1995, p. 212): activities that are determined by, and mediated via, water molecules.

2

Water as a Chaperone

The macromolecular components of cellular systems are either hydrated, are dissolved in water, or they behave in a hydrophobic manner. Indeed, life processes can be defined according to a microbe’s ability to organize materials from its environment to produce or maintain cellular and associated structures (Schro¨dinger, 1944) and, at multiple levels, this process is facilitated by water. Any molecules or ions which enter the cellular system (including those of hydrophobic substances) and are capable of altering the thermodynamic properties of water will affect the structural stability and/or activities of biological macromolecules. Indeed the hydrophobic effect is, arguably, the single most important non-covalent factor in the formation and maintenance of cellular structures: this is true for phospholipid bilayers, as well as for globular proteins in which a delicate balance between entropic and enthalpic effects means that most folded proteins are only marginally stable (20 kJ mol 1) relative to their unfolded state (Fersht, 1985). For DNA, the hydrophobic effect is essential to the stability of the classical, double-helical conformation. This has been demonstrated in studies where synthetic baseanalogues that are unable to hydrogen bond have been substituted for natural bases without any loss of helix stability (Delaney et al., 2003). In the liquid phase, water molecules are constantly in motion, breaking and remaking shared hydrogen bonds. Consequently their entropy is relatively high and anything which reduces this entropy will be energetically unfavorable (Finney, 2004). When non-polar compounds are introduced, water molecules form hydrogen-bonded, cage-like structures around them imposing considerable loss of freedom on those water molecules and consequently reducing the entropy of the system. The energetic cost of this entropy loss, in the range of temperatures consistent with life processes, far exceeds the smaller enthalpic gain so the dissolution of hydrophobic (non-polar) compounds in water is not energetically favorable (Gill and Wadso¨, 1976; Yoshidome et al., 2008; Zielkiewicz, 2008). Since this entropic effect is proportional to the surface area of the non-polar compound (Ho¨finger and Zerbetto, 2005; Wagoner and Baker, 2004), the system will act to minimize this. This principle can be used to describe the internalization of non-polar residues in proteins for the formation of phospholipid bilayers that results in the hydrophobic domains (Chandler, 2005; Dill, 1990; Tanford, 1978) into which lipophilic substances preferentially partition. Biochemical reactions, such as the catabolic breakdown of polyaromatic hydrocarbons, involve contact between two or more organic molecules – in this case an enzyme and a hydrocarbon – that results in modification of one or more covalent bond(s) of the substrate. In this way, cellular processes have a qualitative and potentially irreversible impact on the primary structure of macromolecules and metabolites. By contrast, the behavior of water and hydrophobic molecules in the cell is characterized by cumulatively powerful, but individually weak, interactions (e.g., the hydrophobic effect, hydrogen bonds) that are both reversible and dynamic in nature. Weakly polar (chaotropic) solutes that modify the structure and/or behavior of water

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can induce what is arguably the most potent form of water stress (e.g., Duda et al., 2004; Hallsworth, 1998; Hallsworth et al., 2003, 2007). This said, the tertiary and quaternary structure of macromolecular systems can also be altered by hydrophobic substances such as pesticide residues and organic solvents via mechanisms that are mediated – both mechanically and entropically – by water.1 Direct contact between the polar domains of biological macromolecules and hydrophobic hydrocarbons is minimized by water molecules that collectively act as a mediator, buffer, and lubricant for the movement of hydrophobic substances. Although the term chaperone is typically used to describe proteins that protect and facilitate the formation of protein structures, in terms of these generic activities, water is, in fact, the ultimate chaperone. The cytosol and the extracellular solution constitute two aqueous phases that are notionally separated by the hydrophobic interior of the plasma membrane. This hydrophobic domain effectively behaves as a layer of organic solvent within a water-dominated milieu. For microbes exposed to hydrocarbons, the question arises whether there are hydrophilic-hydrophobic interfaces within the membrane that act as thermodynamic or mechanical barriers regulating the behaviour of hydrocarbon molecules and/or their movement into the cellular system.

3

Cytosol-Lipid Bilayer-Extracellular Solution; an Aqueous Continuum

Hydrophobic compounds that enter the cell from the extracellular environment may have to navigate organic matrices composed of kosmotropic material such as extracellular polysaccharides (EPS), glycoprotein S-layers, or the lipopolysaccharides and peptidoglycans of the microbial cellwall (Sikkema et al., 1995). Such extracellular material can facilitate attachment to hydrophobic surfaces (Liu and Li, 2008; Thompson et al., 2008), and this demonstrates an ability to interact with hydrophobic substances. EPS and other polymeric matrices may thereby intercept the transit of hydrophobic molecules towards the microbial cell. It is the plasma membrane that defines the boundary of the cell, and this is where hydrophobic substances first interact with the living system. At a conceptual level, the lipid bilayer of the plasma membrane represents a three-phase system comprising of an external hydrophilic layer of polar lipid head-groups, that borders the extracellular environment; a second, internal, hydrophilic layer of lipid headgroups that borders the cytosol; and a hydrophobic layer that is sandwiched in-between. Some authors have drawn distinctions between the hydrophobic membrane interior, the glycerol and head-group regions that are in contact with water, and the aqueous milieu outside the membrane (Matubayasi et al., 2008). The membrane therefore presents a series of four notional interfaces across which hydrophobic substances would have to move in order to penetrate and traverse the lipid bilayer; the interfaces between (1) the extracellular solution and the external polar (hydrophilic) layer, (2) this external polar layer and the hydrophobic membrane interior, (3) the hydrophobic interior and the internal polar layer, and (4) the inner membrane surface and the cytosol. There is a substantial quantity of water in the membrane’s hydrophilic layers at any one time that is hydrating the polar head-groups, and the remaining membrane water exists

1

Some of these interactions do not result in metabolic inhibition, for example those between the relatively non-polar enzyme and substrate, naphthalene dehydrogenase and naphthalene, (Kulakov et al., 2006) that give rise to a physiologically valuable response.

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primarily in the bulk-liquid phase (Erilov et al., 2005; Wiener and White, 1992). The hydrophobic interior of the lipid bilayer is heterogeneous in character and is a thermodynamically distinct environment that exhibits similar characteristics to organic solvents. Within the membrane interior the behaviour of the zone closest to the polar head-region most resembles liquid hexadecane, whereas that towards the mid-bilayer has been likened to liquid hexane (Marrink and Berendsen, 1994). Despite the structural and behavioral distinctions between hydrophilic and hydrophobic phases, a close physical analysis of these ‘‘interfaces’’ would prove fruitless because the membrane, in reality, forms a seamless continuum. Some authors have alluded to this by proposing multiple-phase systems with acknowledgement of diffuse boundaries (see Bassolino-Klimas et al., 1994; Marrink and Berendsen, 1994). Water molecules located in the hydrophilic phases of the plasma membrane form a dynamic, but continuous, hydrogenbonded network with those located in the extracellular solution or cytosol. Despite the relatively hydrophobic nature of the membrane interior, even this region is penetrated by some water molecules (see also > Fig. 1; Erilov et al., 2005; Matubayasi et al., 2008). For example,

. Figure 1 Representation of a lipid bilayer showing polar lipid head-groups (bold black lines), gradient of water molecules (blue), gradient of weakly hydrophobic hydrocarbons (orange-red circles), and the location of strongly hydrophobic hydrocarbons (red lines). Modified and reprinted with permission from Marrink and Berendsen (1994). Copyright (1994) American Chemical Society.

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in a model dipalmitoylphosphatidylcholine (DPPC) bilayer used in a 120-ps simulation of water transport through membranes, it was concluded that a small number of water molecules exist in the mid-bilayer at any one time and that they remained for an average of around 10 ps (Marrink and Berendsen, 1994). Substances that accumulate in the membrane interior due to their hydrophobic nature (e.g., benzene) can, nevertheless, be sufficiently polar to form weak hydrogen bonds with water. This association with water molecules may facilitate the passage of such hydrocarbons from outside the cell into the polar head-group region of the plasma membrane. Furthermore, a small percentage of the benzene molecules that enter the hydrophobic interior of the lipid bilayer may act as a vehicle for the transport of (a) water molecule(s) into the membrane interior. Whereas phenol is not hydrophobic enough to partition preferentially into the membrane interior, it can nevertheless traverse the lipid bilayer. The polarity of the hydroxyl group of a phenol molecule may be sufficient to transport water molecules into the interior of the bilayer leaving open the possibility that phenol too can liberate occasional water molecules into the lipophilic region of the membrane. In a number of ways, the plasma membrane forms a continuum with the cytosol and the extracellular solution. Firstly, from a structural viewpoint, lipids, proteins and other molecules in the bilayer form a mechanical continuum across the membrane (see > Fig. 1). Secondly, in terms of temporal events, the kinetic energy of molecules is transmitted across the bilayer, the components of which are in constant motion. Thirdly, the hydrophobic effect on the one hand, and non-covalent interactions such as hydrogen bonds on the other hand, are energetically interconnected and interdependent. Although it is possible to make qualitative distinctions and to envisage discontinuities between the types of forces in different zones of the bilayer (see Marrink et al., 1993), it is irrefutable that the cytosol, lipid bilayer and extracellular solution form a mechanical and energetic continuum, and that the distinction between the hydrophobic and hydrophilic phases of the bilayer is blurred by the continuous motion of water, lipids, and other molecules in the fluid bilayer (Peters et al., 1974; Ritter et al., 2003). It is unfortunate, therefore, that the linguistic expressions required to describe membrane structure (e.g., hydrophobic mid-plane and hydrophobic interior) imply the existence of mechanically separate layers.

4

Behavior of Hydrophobic Substances in the Cellular System

Microbial habitats can present an array of chemically diverse hydrocarbons to the cell (for examples see > Vol. 1, Part 3 and > Vol. 2, Part 14). In oil-polluted environments these include low-molecular weight, relatively soluble aromatic compounds such as benzene, as well as long-chain aliphatic, strongly hydrophobic substances such as hexadecane (Sikkema et al., 1995). Given the limited solubility of hydrophobic hydrocarbons in water, their concentration in the aqueous extracellular milieu can be orders of magnitude lower than that of common hydrophilic solutes such as Na+, Cl– and NH4+ ions, urea, glucose, and sucrose. Nevertheless, bioavailable hydrophobic molecules can be even more potent as cellular stressors than osmotically active or chaotropic substances when compared on a molar basis (Bhaganna and Hallsworth, unpublished data; Hallsworth et al., 2003, 2007; Vermue¨ et al., 1993). Water in the hydrophilic regions of the plasma membrane is typically more structured (see Marrink and Berendsen, 1994; Marrink et al., 1993) than that outside the cell due, we believe, to the kosmotropic activity of polar lipid head-groups in the bilayer: the increased order arises

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from increased-affinity hydrogen-bonding between water molecules and polar lipid headgroups. In addition, we hypothesize that the water adjacent to the hydrophobic mid-plane can be ordered via the same process that results in cage-like structures of water molecules around dissolved hydrocarbons; in this case increased order is associated with the lack of hydrogenbonding between water molecules and the hydrophobic region (see above). In consequence, the diffusion rate of molecules in the polar head-region is reduced by an order of magnitude (Marrink and Berendsen, 1994) such that the majority of dissolved hydrocarbon molecules – and indeed those of any other solute – tend to remain outside the cell membrane unless taken up by a transport protein. We speculate, therefore, that the concentration of hydrophobic compounds in the polar region of the plasma membrane is usually less than that outside the cell. Nevertheless, a proportion of the dissolved hydrocarbon molecules that come into contact with the plasma membrane does enter the hydrophilic phase of the bilayer and two mechanisms have been identified by which this occurs: the transient channel and the solubility-diffusion mechanism (Gupta et al., 2008). Firstly, the molecular motion of bilayer molecules can lead to the formation of transient channels across the membrane, through which hydrocarbon molecules can move. Secondly, short-lived cavities that appear in the polar, hydrophilic region(s) can facilitate entry of hydrophobic molecules into the membrane, following which these molecules may move within the bilayer via other voids that momentarily appear. The formation and frequency of these voids are likely to increase in the proximity of the membrane interior; indeed motion of hydrophobic substances becomes more free in proportion to their proximity to the mid-bilayer (Bassolino-Klimas et al., 1994). The spasmodic motions of hydrophobic substances that move through the polar-head region via spontaneous voidformation may resemble those of the occasional water molecules that diffuse into the hydrophobic bilayer interior. As explained by Marrink and Berendsen (1994), the ‘‘diffusional jumps’’ of a water molecule become greater the closer it gets to the mid-plane. We hypothesize that some hydrocarbon-degrading microbes may respond to hydrophobic compounds by modifying their plasma membrane structure in such a way that facilitates the spontaneous entry of substrate molecules into the cell. The partition coefficients for hydrocarbons in two-phase organic-solvent:water systems (commonly octanol:water) can be used to quantify the hydrophobicity of the former. Alternatively, water:lipid-membrane vesicles can be used for this purpose (Efremov et al., 2007; Kwon et al., 2006). Data obtained from reductionist systems may not, however, be pertinent to the cellular system because the partitioning of hydrocarbons into the hydrophobic interior of biological membranes is determined – in part – by the composition of the membrane (lipids, proteins etc.; Sikkema et al., 1994), and is spatially heterogenous within the bilayer. Weakly hydrophobic substances, such as benzene, will accumulate throughout the lipid acyl-chain region (see > Fig. 1; Marqusee and Dill, 1986; Norman and Nymeyer, 2006) whereas strongly hydrophobic compounds are concentrated towards the centre of the bilayer (see > Fig. 1; Xiang and Anderson, 1994). For strongly hydrophobic substances there is, therefore, a concentration gradient of hydrophobic molecules across the plasma membrane in the converse direction to that of water molecules (see > Fig. 1; Marrink and Berendsen, 1994). Towards the centre of the membrane there is a disorder gradient for both membrane-lipid molecules, and for hydrocarbons that have entered the bilayer, with disorder increasing towards the mid-plane (Dill and Flory, 1980). Both the strength of the hydrophobic effect and the mechanical consequences of this disorder gradient may act as driving forces to enhance accumulation of hydrophobic compounds towards the bilayer mid-plane (Xiang and Anderson, 1994). Generally, there is a limit to the quantity of hydrophobic molecules that the plasma

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membrane can accommodate, beyond which bilayer stability becomes compromised. One study of a range of gram-negative and gram-positive bacteria found that the critical concentrations of organic solvents that could dissolve in the plasma membrane did not vary between microbial species (Vermue¨ et al., 1993). For small cyclic hydrocarbons such as benzene and toluene as this concentration limit (i.e., approximately one hydrocarbon molecule per two phospholipid molecules) is approached, the lipid bilayer becomes increasingly disordered and the membrane becomes more porous (Sikkema et al., 1994). This affects cell turgor, energy generation, solute transport, signal transduction and other critical cellular functions (Sikkema et al., 1995). Both long-chain and low-molecular weight molecules weaken interactions between membrane lipids and can, at high concentrations, cause expansion of the membrane. Intermediate molecular-weight hydrophobic substances such as Lindane (d-hexachlorocyclohexane, mw 290.85) have been found to disorder liposome membranes and, even at subsaturated concentrations, can induce cell lysis in vivo (Sikkema et al., 1994). This said, long-chain and branched hydrocarbons spanning both monolayers can have an additional stabilizing effect on membrane structure that may even result in a net increase in membrane lipid order (see Sikkema et al., 1995). At multiple levels, the chemistry and functionality of proteins that are located within a lipid bilayer are determined by the hydrophobic effect (see Lomize et al., 2004). For instance, the alpha-helices of membrane proteins may interact using hydrophobic or polar amino acids. In the hydrophobic environment of the membrane interior there will be a thermodynamic tendency towards the minimisation of the surface area of polar regions, thus promoting association). Furthermore, polar interactions in the interior of all protein molecules (again, energetically driven by the hydrophobic effect) can confer structural and functional stability (Lomize et al. 2004; Zhang and Lazaridis 2006). The location of polar and/or hydrophobic domains of proteins within a lipid bilayer, as well as the movement of hydrophobic molecules (or, indeed, polar solutes) into the protein molecule, can therefore determine structural stability of proteins, membranes and other cellular macromolecules (Gratkowski et al. 2001; Lear et al. 2003; Vijayan et al. 2005; North et al. 2006). Individual hydrocarbon molecules will remain within the hydrophobic interior of the plasma membrane for a mean period of time that depends on factors such as hydrocarbon concentration, membrane composition, and temperature (see Bassolino-Klimas et al., 1994; Sikkema et al., 1995). We speculate that the residence time for hydrocarbons in the mid-plane is also dependent on the degree of water structure in the polar head-regions of the bilayer. A proportion of these molecules will re-dissolve in either the outer or inner hydrophilic regions of the bilayer, from where they can diffuse into the extracellular environment or the cytosol, respectively (see > Fig. 2). Hydrocarbons dissolved in the cytoplasm are capable of interacting with all cellular macromolecules that have hydrophobic properties. For example, benzene and other hydrocarbons can partition into the hydrophobic interior of cytosolic proteins (Zhang and Johansson, 2005). Hydrophobic cavities in proteins have a finite capacity to accommodate hydrocarbon molecules: those of some proteins are known to be able to accommodate several benzene molecules (see Tanford, 1973, pp. 132–133; Zhang and Johansson, 2005). For hydrophobic hydrocarbons in ribosomes, nucleic acids, or cytosolic- or membrane-bound proteins we speculate that there is a dynamic movement of hydrocarbon molecules into and out of the macromolecular structure. When a microbial cell is exposed to an external source of hydrophobic molecules, there is a net influx of the compound into the plasma membrane and the primary stress mechanism typically involves an alteration of lipid-lipid and lipid-protein interactions (Sikkema et al., 1995). However, hydrocarbon molecules that shuttle between

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. Figure 2 Illustration of the way in which benzene molecules () may behave in relation to a microbial cell situated in an aqueous environment close to a source of crude oil (solid black region) showing: (a) symmetrical exposure of the cell to dissolved benzene, (b) asymmetrical exposure of the plasma membrane to benzene coupled with motion of benzene (see grey arrows) within the membrane, (c) asymmetrical exposure of the plasma membrane to benzene coupled with asymmetrical entry of benzene into the cytosol.

cellular macromolecules and/or cascade from one macromolecular system to another via the cytosol will ultimately impact an indefinite number of cellular structures and processes. The interaction between hydrophobic substances and the cellular system is undoubtedly determined by numerous physicochemical parameters. However, hydrocarbons may also act as toxicants that inhibit metabolism via specific modes-of-action that target, for example, a single enzyme in a specific biochemical pathway (see Trinci and Ryley, 1984). Not only is the mechanism of inhibition highly specific, but each type of toxicity takes place at a specific intracellular site: in this way the inhibitory activity of a toxicant is a metabolically asymmetrical phenomenon. By contrast, for hydrocarbons that act as stressors with generic modes-of-action, the question arises whether the cellular stress induced is always exerted in a symmetrical manner.

5

Do Hydrocarbons Induce Asymmetrical or Symmetrical Stresses?

Microbial cells in natural habitats can be exposed to hydrophobic solutes in homogeneous solutions (e.g., subsaturated concentrations of benzene; see > Fig. 2a), or hydrophobic substances emanating from single-point sources such as small crystals. In other words, hydrocarbon sources can be distributed either symmetrically or asymmetrically in the extracellular environment. For example, hydrocarbonoclastic cells may aggregate around a droplet of crude oil floating on the surface of seawater and are therefore asymmetrically exposed to the oil. A single-point crystalline source presents cells with a gradient of dissolved hydrocarbon, suggesting that the plasma membrane closest to the source will be directly affected by the impact of the stressor, and that cells are therefore asymmetrically stressed (see > Fig. 2c). However we speculate that, in reality, hydrocarbons that enter the external polar-region of the plasma membrane will not come into direct contact with the lipid polar-head groups because of the hydration shells of the latter. We also hypothesize that, upon penetration into the hydrophobic

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interior of the plasma membrane, hydrocarbon molecules will rapidly distribute around the entire membrane (see > Fig. 2b). For low-molecular weight, weakly hydrophobic hydrocarbons such as benzene, this process is likely to occur within seconds or – at the most – several minutes given that lipid molecules introduced at a single point in the bilayer have been shown to redistribute within 40 min (see Peters et al., 1974). There is some evidence that hydrocarbons accumulate in a slightly asymmetric manner in the mid-layer, towards the inner monolayer (Marqusee and Dill, 1986) and, for this reason, the internal (cytosol-) side of the membrane may become disproportionately stressed. The physicochemical properties of both the lipid bilayer and the hydrocarbon(s) will determine the rate at which this redistribution takes place. The distribution of hydrophobic substances in the extracellular environment, combined with the speed at which hydrophobic molecules disperse throughout the cell, with determine the whether the cell is stressed in a way that is spatially symmetrical (see > Fig. 2). This said, the plasma membrane has a spatially heterogeneous distribution of lipid-, protein- and other molecules so it could be argued that the distribution of hydrocarbon molecules and, by implication, the stress that they impose on the membrane system is never completely symmetrical in nature. Conceptually, cellular stress can be categorized as symmetrical or asymmetrical according to diverse criteria; e.g., spatial, temporal or chemical parameters. For cells that are exposed to complex hydrocarbon mixtures such as crude oil, the plasma membrane will simultaneously receive an array of oil-derived molecules. However the molecular kinetics involved in the dissolution and diffusion for diverse components of crude oil will result in temporal fluctuations in the composition of hydrocarbon mixtures to which the plasma membrane is exposed, leading to temporal asymmetry of the hydrocarbon-induced stress. Diverse hydrocarbons in the plasma membrane (or elsewhere in the cellular system) can exhibit qualitative differences in their modesof-action as cellular stressors, and in this way a macromolecular system can be exposed to chemically asymmetrical challenges. That microbial cells can become asymmetrically stressed by hydrophobic substances has a number of biological implications. For example, it is intriguing to question whether there is a correspondingly asymmetrical cellular-stress response. Conversely, if a microbial cell does exhibit an asymmetrical stress response, this may be used for diagnostic purposes to determine whether the cellular system had been inhibited via an asymmetric stress mechanism. For any specific hydrocarbon, there is a degree of freedom in the interaction between the following factors: the extent to which it penetrates the cell, its inhibitory effects on the functionality of macromolecular systems, and the consequential cellular stress response/adaptation. The kinetics of molecular motion associated with the diffusion and dispersion of hydrocarbon molecules, or to the instantaneous movement of low molecular-weight hydrocarbons around the mid-plane of the plasma membrane (> Fig. 2b), may take place within a timescale that is too short for a biological (genomic) response to have been triggered.2 In other words, we suggest that the stress exerted against the macromolecular system can sometimes become diluted, or even eliminated, before sufficient time elapses for the cell to sense and respond. Furthermore, whereas hydrophobic stressors disorder membrane, protein, and nucleic-acid structures; other substances or environmental conditions may counter this effect. A reduction in temperature or an increase in the concentration of kosmotropic solutes such as NaCl, sucrose, trehalose, or betaine 2

This said, a more rapid response may occur at a phenotypic level; for example the cis-trans unsaturated fatty-acid isomerase of Pseudomonas putida. This enzyme is expressed constitutively at all stages of the growth cycle but appears only to be active only when the cell is challenged by a stressor, such as toluene (Bernal et al., 2007; Junker and Ramos, 1999).

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will act to re-order cellular macromolecules and membranes. The quantitative and qualitative impacts of hydrophobic stressors on the cellular system are determined by the net effect of the cellular phenotype (and the degree to which it can respond and adapt), other dissolved substances, and environmental conditions.

6

Microbial Cells in Hydrophobic Environments: Situational-Functional Considerations

Both aqueous and terrestrial habitats in Nature can contain non-dissolved sources of hydrophobic materials such as droplets of crude oil in seawater, or crystals of Lindane in soils (> Fig. 2; Thomas et al., 1996). Similarly, the microbial environments of industrial systems may contain undissolved forms of hydrocarbon, such as the toluene in some two-phase biocatalysis systems (see Heipieper et al., 2007 [> Vol. 3, Part 5]). Nevertheless, in the context of the vital regions of a cell, i.e., the cytoplasm and plasma membrane, it is questionable whether cellular macromolecules are indeed in direct contact with extracellular bodies of hydrophobic material. Microbial cells can produce EPS, cell-wall material and biofilms that act as mechanical and physicochemical buffers that prevent contact between the plasma membrane and hydrophobic bodies (for an example, see Battin et al., 2007). In the case of a naked cell that is not protected in these ways, the outer surface of the plasma membrane is polar and the lipid headgroups are surrounded by water and will therefore be mechanically, and energetically, protected from contact with undissolved hydrocarbons. As pointed out by Sikkema et al. (1995), direct contact between hydrophobic substances and the interior of the plasma membrane is prevented by the hydrophilic parts of the outer membrane, or otherwise by the cell wall. Extracellular structures (both biotic and geochemical) such as cell-wall material, EPS, and clay particles, can sequester and retain, and may also release, hydrophobic molecules (see Abbasnezhad et al., 2008; Strevett and Chen, 2003). It is plausible that cells utilize these activities by synthesizing or organizing structures for this purpose. In a study of clay micelles, bacteria formed ‘‘hutches’’ around their cells, that appeared to sequester and supply hydrophobic compounds that could then be catabolized cell (Lu¨nsdorf et al., 2000). The implication of, what appears to be, a clay-micelle shuttle for hydrophobic compounds is that the cells receive a controlled stream of compounds that can be used as a consistent supply of nutrients whilst avoiding unlimited levels of cellular stress. Microbes that degrade hydrocarbons, especially those that synthesise biosurfactants to enhance the bioavailability [> Vol. 2, Part 6], must adapt to the stress induced by these hydrophobic substances. Although biosurfactants/bioemulsifiers are often considered to act simply as dispersants, they probably coat their hydrophobic targets, as well as the producing cell. Alcanivorax borkumensis, for example, produces a glucose-lipid surfactant that has distinct chemical forms: a cell-attached and an excreted version (Yakimov et al., 1998). Interactions between cell and substrate may well be surfactant:surfactant molecule rather than hydrocarbon:plasma membrane interactions. Generally, surfactant molecules tend to have a disordering or chaotropic effect on lipid bilayers. Whereas the utilization of hydrocarbons for microbial growth can enhance species competitiveness (Head et al., 2006), we speculate that the ecological strategy of degrading hydrocarbons concomitantly imposes a metabolic cost on the cell. Strongly hydrophobic substances, with a log Poctanol-water 5, are typically so insoluble in water that microbial cells have little exposure to them (Usami et al., 2003). Weakly hydrophobic

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compounds such as benzene are relatively soluble in water and are accordingly bioavailable (Sikkema et al., 1995). Generally speaking, substances that are strongly hydrophobic are more disruptive to cellular structures and are therefore highly inhibitory, or even lethal, at lower concentrations. There is, therefore, an interaction between the hydrophobicity of hydrocarbon stressors, their bioavailability, and the level of cellular stress induced per mole. On balance, the most stressful compounds for microbial cells have a log Poctanol-water in the range 2–4 (Inoue and Horikoshi, 1991). Some microbes, such as the bacterium P. putida, can catabolize a vast number of hydrophobic substances (see Timmis, 2002). Species that have evolved to utilize hydrocarbons are at the same time metabolic specialists and highly versatile, enhancing their ecological competitiveness, both by harnessing a wide array of nutrients, and by reducing cellular stress via the catabolism and detoxification of potentially lethal substances. In this way catabolism of hydrophobic substances can be seen as a mechanism that has evolved to evade, or at least minimize, cellular stress (see Dominguez-Cuevas et al., 2006); a strategy most likely to succeed when stressor molecules are in limited supply. There are diverse types of inclusion bodies found in microbial cells which contain, or are composed of, hydrophobic substances. Some microbes may form inclusion bodies which separate hydrophobic substances from the surrounding cytosol and thereby protect cellular macromolecules from hydrocarbon-induced stress (Kim et al., 2002). Bacterial cells have been shown to evade or minimize damage by stressors such as toluene by producing membrane vesicles that contain the hydrocarbon and can be expelled into the extracellular environment (Kobayashi et al., 2000), whereas other cells that are highly tolerant to hydrophobic substances use efflux pumps to remove stressors from the cell (Kieboom et al., 1998; Ramos et al., 2002). It is well-known that the physicochemical properties of specific proteins can make them more or less resistant to structural changes induced by hydrophobic solutes (Dohnal et al., 2001). Some halophilic bacteria and Archaea that accumulate ions in the cytosol as osmolytes have evolved to synthesise proteins that function optimally under extremely saline conditions (Kushner, 1978). It is not implausible, therefore, that organic-solvent tolerant-microbes or hydrophobic-stressor tolerant-microbes exist that have been selected for their ability produce proteins that are structurally stable under these conditions.

7

Research Needs

While there has been a considerable research effort to understand the interactions between hydrophobic substances, water, and lipids and/or proteins in vitro, there has been relatively little work to elucidate how hydrophobic substances interact with the global cellular system as stressors. Compounds with a log Poctanol-water 2 have traditionally been perceived as toxicants and not stressors, but these substances undoubtedly perturb macromolecular structures and interactions in a non-specific manner, and the generic stress mechanisms in vivo and corresponding cellular responses have not yet been fully elucidated. Furthermore, the commonality and distinctions between chaotrope-induced stress mechanisms and responses (see Hallsworth et al., 2003, 2007) and those induced by hydrophobic substances have not yet been explored. One other outstanding question is whether cells can be affected by hydrocarboninduced stresses in an asymmetrical fashion, or whether the physics of molecular motion is always so rapid that the cell’s biological machinery does not respond within the timeframe required for the physical parameters to entropically redistribute in a symmetrical fashion.

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Hydrophilic polymers that are kosmotropic, such as those that make up EPS, withhold water in a matrix (see Brown, 1990, p. 35). It is not implausible, therefore, that the ordering effect (see Eisenberg and Kauzmann, 1969) of these macromolecules on the hydrogen-bonded network of water molecules creates, or at least contributes to, a barrier-effect around the cell by enhancing water-water interactions, thereby reducing the transit of hydrophobic molecules towards the cell. Whereas chaotropic molecules, such as ethanol, have apparently moulded the ecological behavior of some microbial species (see Hallsworth, 1998) there is currently a large knowledge gap in relation to the manner in which the stresses induced by hydrophobic substances have determined, and how they continue to impact on the evolution of microbial species and their macromolecules. The combined implications of the bioavailability of hydrophobic substances, their value as nutrient sources, the potential stress effects of biosurfactant molecules, and the potency of hydrophobic substances as stressors have not yet been elucidated in the context of the biology and ecology of the microbial cell. It is intriguing to question how the stress tolerances and the specific stress-responses of diverse microbial species determine both their ecological strategies and evolutionary trajectories.

Acknowledgments We are grateful for thought-provoking discussions with Giuseppe Albano (Edinburgh University, UK), Prashanth Bhaganna and Kalpa D. Gupta (Queen’s University Belfast, Northern Ireland), Ananda Hillis (University of Ulster, UK), Allen Y. Mswaka (University of Harare, Zimbabwe), Mary Palfreyman (Outwood Grange College, UK), Harald J. Ruijssenaars (TNO Quality of Life, The Netherlands), Kenneth N. Timmis (HZI, Germany) and Graham J. C. Underwood (University of Essex, UK). Work on this article was funded by the Kluyver Centre for Genomics of Industrial Fermentation (The Netherlands), EU Fifth-Framework contract QLK3-CT-200201933 (LINDANE), Biotechnology and Biological Sciences Research Council (BBSRC, UK) and Natural Environment Research Council (NERC, UK).

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44 Matrix-Hydrophobic Compound Interactions H. Harms1,* . L. Y. Wick1 . K. E. C. Smith2 1 Department of Environmental Microbiology, UFZ, Helmholtz Centre for Environmental Research, Leipzig, Germany *[email protected] 2 Department of Environmental Chemistry and Microbiology, The National Environmental Research Institute (NERI), Roskilde, Germany 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1468 2 Interactions of Non-Aqueous Phase Liquids with Solid Matrices . . . . . . . . . . . . . . . . . 1468 3 Phenomena of Molecular Sorption to Solid Matrices . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1469 4 Partitioning of Hydrocarbons between the Aqueous and the NAPL-Phase . . . . . . . . 1470 5 Sorption to Mobile Sorbents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1473 6 Contaminant Aging and Release Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1475 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1476

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_100, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Fate and transport of hydrophobic organic compounds (HOC) are strongly influenced by their interactions with environmental matrices. These interactions can be grouped into those of non-aqueous phase liquids (NAPL), i.e., spreading of oil on solid surfaces and its movement in porous media and those of predominantly water-dissolved molecules which may sorb onto solids or partition into organic matter or NAPL phases. All these types of sequestration phenomena lead to reduced contact of water-borne organisms with the HOC. In contrast, sorption to mobile matrices may mobilize HOC and thus increase bioavailability and toxicological risk.

1

Introduction

Microbial degradation of hydrocarbons requires physical contact between the organisms and the chemicals. The same holds true for toxic effects of hydrocarbon compounds on biota. Environmental matrices constituted of inorganic and organic materials interfere with this contact. They may constitute barriers, which at the macroscopic scale keep reservoirs of the chemical separate from habitats of microbial communities. At the molecular scale they may hinder individual hydrocarbon molecules from diffusing to soil or sediment pores colonized by microbes. Biodegradation becomes limited by the separation and when the hydrocarbon serves as the growth substrate, its physical confinement limits the build-up and maintenance of microbial biomass. In ecological terms, matrices may affect the carrying capacity of a habitat via their impact on substrate accessibility. On the other hand, physical confinement may drastically reduce the toxic risk arising from environmental chemicals. This chapter classifies matrix-compound interactions and describes their effects on hydrocarbon microbiology.

2

Interactions of Non-Aqueous Phase Liquids with Solid Matrices

Interactions of NAPLs with solids are of importance wherever oil is present in porous media. This is the case for instance in oil reservoirs. Here the interest in extracting the oil has motivated much research into the possibility to pump the oil out or, if pumping fails, to push residuals out of the porous matrix by injecting gases or aqueous solutions (e.g., brine), sometimes in combination with selective blocking of alternate flow paths, thermal treatment (e.g., steam injection) or physicochemical stimulation of oil movement using detergents (Banat, 1995). A second field of environmental concern is the behavior of spilled fuels, coal tar or other oily masses in soil, aquifer sediment, fractured rock, on beaches and in the sediments of rivers, lakes and the ocean. A detailed description of the physics of the residence behavior and movement of oil in these phases would go beyond the purpose of this chapter, but some factors of influence shall be mentioned here. Oil masses of lower viscosity move more readily in porous media. The viscosity depends on the chemical composition of the oil and can be reduced by an increase in temperature. The injection of gases in enhanced oil recovery also reduces the viscosity of oil as some of the gas dissolves in the oil, a phenomenon called oil swelling (McInerney et al., 2007). Emulsification of oils with water can influence the viscosity in both ways depending on the oil-water ratio, i.e., if the oil is the continuous phase or the dispersed phase in the emulsion, but also depending on other factors such as the size of

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the droplets of the dispersed phase. An illustration of this effect is the deviation of the viscosity of the water in oil emulsion that we call margarine from the weighted mean viscosity of its components. In the environment quasi-solid emulsions of hydrocarbons and water have been observed as viscous interfacial films around aged tar globules (Nelson et al., 1996). Emulsifiers are surface-active substances capable of stabilizing emulsions by accumulating at the water oil interfaces. Both the chemical composition of the solid material and the pore diameter of the solid matrix influence the oil movement via capillary forces. If the oil has a tendency to spread on the solid or, in other words, to form a contact angle with the solid surface below 90
, it will be retained better in pores of smaller diameters. The opposite will be the case if the oil forms a contact angle above 90
with the solid surface. In this case capillary forces will retain the oil better in larger pores. This behavior is described by the Young-Laplace equation (Mozes et al., 1991). Capillary forces can be influenced by surfactants of chemical or biological origin leading to improved movement of the oil in the porous medium.

3

Phenomena of Molecular Sorption to Solid Matrices

The total amount of a specific compound can be conceptually divided into three pools, i.e., the irreversibly-bound, the reversibly-bound, and the freely dissolved pool (> Fig. 1). The mechanistic interpretation of the macroscopically observed sorption behavior of hydrophobic organic chemicals in soils and sediments has been an issue of much debate in the last decades. It is therefore highly appreciable that in 1997 ten of the leading experts in the field, among them proponents of contrasting views, published jointly an article that categorizes the various mechanisms of HOC sorption to geosorbents (Luthy et al., 1997). These authors distinguish

. Figure 1 The total amount of a compound in an environmental system can be divided conceptually into the freely dissolved, the reversibly- and the irreversibly-bound pools. From the perspective of microbial hydrocarbon degradation, the kinetics of the release of HOC into the water phase, i.e., into the freely dissolved pool, is of primary importance. Soft organic matter and NAPL-absorbed HOC as well as surface-adsorbed HOC appear to be more readily bioaccessible than quasi irreversibly-bound HOC, ‘‘stuck’’ in hard organic matter and micropores.

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five microscopic sorption mechanisms of HOC in geosorbents: (1) absorption into amorphous or ‘‘soft’’ natural organic matter or NAPL, (2) absorption into condensed or ‘‘hard’’ organic polymeric matter or combustion residues such as soot, (3) adsorption onto water-wet organic surfaces such as soot, (4) adsorption onto exposed water-wet mineral surfaces, and (5) adsorption into microvoids or microporous minerals. An important conclusion was that none of these sorption mechanisms is likely to occur exclusively in natural geomaterials and that complex sorption and desorption behaviors in terms of equilibria and kinetics are explainable as overall effects of varying contributions of these mechanisms. These mechanisms were then examined for their impact on the behavior of HOC in terms of the linearity of sorption isotherms, the possibility of competition of sorbates for sorption sites, sorption kinetics and, if there is desorption hysteresis, selectivity for steric features of sorbents, the ease at which the HOC can be extracted and other features of less importance for hydrocarbon microbiology. Absorption into soft organic matter and NAPL ((1) in the above list) as well as both types of adsorption onto exposed surfaces (3, 4) were identified to be fast and readily reverted by solvent extraction, whereas absorption into hard organic matter (2) and adsorption in microvoids (5) were characterized as slow, difficult to revert by solvent extraction and showing desorption hysteresis. From the perspective of microbial hydrocarbon degradation, the kinetics of the release of HOC into the water phase is of primary importance. Soft organic matter and NAPL-absorbed HOC as well as surface-adsorbed HOC appear to be more readily bioaccessible than HOC ‘‘stuck’’ in hard organic matter and micropores. The case of pore-obstructed HOC illustrates that there is also a geometric aspect of bioaccessibility (in this case the exclusion of microbes from micropores that spatially separates the HOC source from its biological sink) next to the influence of chemical interaction. This is due to the dynamic nature of microbial degradation and the fact that rates of mass transfer depend largely on the distances that need to be bridged (Bosma et al., 1997).

4

Partitioning of Hydrocarbons between the Aqueous and the NAPL-Phase

NAPLs are varied with respect to both the environmental compartment in which they are found, for example the open sea, sediment or soil, but also to their physical and chemical characteristics. Typical examples of environmentally relevant NAPLs include crude oil, its various refinement products and anthropogenic wastes such as the aromatic rich coal-tars contaminating groundwater at industrial sites. It is interesting to note that there are natural oil seeps in the marine and terrestrial environment. Hence, the interplay between microorganisms and certain types of NAPLs is not only a recent phenomenon, but there has been sufficient time to evolve strategies for an increased exploitation of this rich hydrocarbon resource (Head et al., 2006). NAPL hydrocarbons represent a potential source of carbon and energy in a form that is difficult to exploit, since it appears that hydrocarbons need to exist in the dissolved state before they can be taken up (Volkering et al., 1992). Thermodynamic considerations indicate that even at equilibrium, i.e., the maximum possible dissolved aqueous concentrations that can be attained, hydrocarbons preferentially remain in the NAPL with only low concentrations being reached in the aqueous phase (Efroymonson and Alexander, 1995). Furthermore, in nonequilibrium situations these same properties mean that the mass transfer between the NAPL and the aqueous phase is usually slow (Schluep et al., 2002). Therefore, it is usually the case

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that NAPL hydrocarbons have a low bioavailability and biodegradation can be mass transfer limited (e.g., Ramaswami et al., 1997). Note though, that although this low bioavailability is not ideal for a degrading microorganism, it also means that for non-degrading organisms exposure to potentially toxic NAPL phase hydrocarbons is reduced. NAPLs are typically composed of many classes of hydrocarbons in combination with other non-hydrocarbon compounds that are often unresolved, e.g., crude oil (Marshall and Rodgers, 2004). It is therefore not usually the case that a single micro-organism has the metabolic ability to degrade the full range of components, and therefore a range of micro-organisms are involved in the biodegradation process of NAPL hydrocarbons (Head et al., 2006). Therefore, in the field biodegradation is a complex interplay between bioavailability, toxicity and microbial ecology. The interface between NAPL to the aqueous phase can be considered as being composed of two barriers to mass transfer aligned in series (Schwarzenbach et al., 2003). Adjacent NAPL and aqueous compartments, each with homogenous bulk hydrocarbon concentrations, are separated by the NAPL:water interface. In each phase, thin unstirred boundary layers (BL) exist as depicted below in > Fig. 2. Within these, transport of the hydrocarbons takes place by the relatively slow process of molecular diffusion. Therefore, the BLs (in most cases the BL on the aqueous side; Ghoshal and Luthy, 1996) have the determining role in the overall mass flux into the bioavailable dissolved phase. Special cases where the main resistance to mass transfer occurs in the NAPL phase are discussed at the end of this section.

. Figure 2 Chemical activity profile (- - - - -) of a hypothetical hydrocarbon across the NAPL: aqueous interface under various environmental conditions. The resultant effect on the magnitude of the hydrocarbon mass flux is shown at the bottom by a positive or negative symbol.

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The hydrocarbon mass flux can be defined as the mass from the NAPL phase entering the aqueous phase where the degrading micro-organisms are present over a given period of time. This can be described in general terms as follows Mass flux ¼ Area  Transfer velocity  Driving force

ð1Þ

The role of the interface area is obvious, the larger this is the more surface there is for exchange. The transfer velocity can be further rationalized as being composed of the ratio between the diffusivity of the hydrocarbon and the thickness of the BL (Schwarzenbach et al., 2003). This is also intuitively obvious, a higher diffusivity of a molecule is indicative of more rapid motion and a thinner BL will be more quickly traversed. The driving force is determined by the gradient in chemical activity between the bulk NAPL and aqueous compartments. Molecular diffusion occurs from regions of high to low chemical activity (Reichenberg and Mayer, 2006), and the greater the difference the greater the hydrocarbon flux. Chemical activity, which might be viewed as the ‘‘temperature’’ of a chemical, is a function of the properties of the compound and the environmental matrix in which it is found. Therefore, a hydrocarbon can have the same chemical activity in two phases but very different concentrations. A well known illustration of this is the equilibrium octanol:water partition coefficient. At equilibrium, a hydrocarbon compound has the same chemical activity in the water and octanol phases, despite the very different concentrations. However, the usual practice is to measure mass concentrations in the NAPL and aqueous phases. The driving force is then computed from these concentrations in conjunction with experimentally determined partition coefficients (e.g., Schluep et al., 2002). > Equation 1 indicates that the magnitude of the mass flux is related to a number of factors including the NAPL:aqueous interface area, the speed of transfer across the BLs and the driving force between the NAPL and aqueous compartments. Therefore, a change in any one of the above can result in an increase or decrease in the mass flux of hydrocarbon into the aqueous phase, and thus have a knock-on effect on bioavailability and biodegradation. This is shown schematically in > Fig. 2, where different scenarios have been depicted in terms of the above three factors. Here it can be seen, a range of abiotic processes can have a potential big impact on the mass transfer of hydrocarbons into the bioavailable aqueous phase (> Fig. 2A–D). Furthermore, these can occur simultaneously and may vary with respect to one another over time. These have been summarized in > Table 1. Some of these changes can have other beneficial side-effects with respect to microbial growth, in addition to any enhancement in the mass transfer. For example increased mixing might also lead to improved aeration and prevent the formation of nutrient-depleted patches. Changes in temperature might lead to an increase (or decrease) in growth depending on the degrader. In specific situations, the resistance to diffusive mass transfer on the NAPL side can become significant (> Fig. 2). Such scenarios include the mass transfer of more soluble hydrocarbons (Schluep et al., 2002), highly viscous NAPLs (Ortiz et al., 1999) or situations where there is a weathering of the surface layers of multicomponent NAPL mixtures leading to the formation of more impermeable surface skins. The latter is believed to be the result of viscous NAPL:water emulsions forming at the surface rather than because of changes in the composition due to preferential dissolution of the more soluble components (Nelson et al., 1996). In terms of microbial degradation of NAPL hydrocarbons such cases are significant for two reasons. Firstly there can be a reduction in the mass transfer into the aqueous phase (Luthy et al., 1993). Secondly, the above environmental processes affecting the mass transfer through the aqueous boundary layer no longer have the same role. An obvious example

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. Table 1 Overview of the physical, environmental and chemical factors potentially influencing the mass transfer of a hydrocarbon from the NAPL into the aqueous phase. The letters in brackets correspond to the scenarios depicted in > Fig. 2. The literature references are studies where the changes described have been shown to influence the microbial degradation of NAPL phase hydrocarbons Category Physical

Description NAPL architecture and amount e.g., film, droplets, micropores

Environmental Hydrodynamic mixing

Parameter affected Surface area [A] Break-up of NAPL [A] BL thickness [B]

Temperature

Diffusivity [C] Chemical activity gradient [D]

Chemical

Sorption in the aqueous phase

Chemical activity gradient [D]

Hydrocarbon concentration

Chemical activity gradient [D]

Hydrocarbon properties

Diffusivity [C] Chemical activity gradient [D]

NAPL properties

Diffusivity [C] Chemical activity gradient [D] Wetting properties [A]

Weathering of multicomponent NAPLs

Formation of impermeable skins [E]

NAPL side resistance

Diffusivity and chemical activity [E] Gradient

involves changes in the degree of hydrodynamic mixing, which will no longer have the effect of increasing or decreasing the mass transfer. From the point of view of the degrading micro-organism, there is little that can be done to influence such abiotic processes and thus they can only react passively to any resulting changes in the mass flux and bioavailability. An interesting example is that of sorption, which had the consequence of lowering the dissolved phase activity and thus maintaining a high diffusive gradient as illustrated in > Fig. 2D. Here there is a feedback between biotic growth of the micro-organism due to degradation and the abiotic sorption process. A bacterial population actively degrading a certain NAPL component will increase in biomass, i.e., an increase in sorption capacity, thus potentially enhancing the mass transfer of other NAPL hydrocarbons.

5

Sorption to Mobile Sorbents

Various types of hydrocarbon interactions with the geosorbents present in soils and sediments have been considered above. These sorbents can in the main be considered as relatively immobile, with only small fractions being slowly displaced by processes such as bioturbation or erosion by wind and water. Therefore, for microorganisms to colonise such sorbents, they need to initially be brought into contact with and subsequently attach to the surface. Some

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sorbing phases, though, are much more mobile, moving with the aqueous phase. This is a diverse category, encompassing everything from suspended inorganic minerals, particulate and dissolved organic matter, living bacteria- and phytoplankton to biosurfactants. Of course there is no defined cut-off between what constitutes a mobile and immobile sorbing phase, and furthermore, the significance they have in terms of the total hydrocarbon sorption depends on the environment in which they are found. For example in soils and sediment most of the hydrocarbons are primarily associated with larger sized and stationary particulate material (e.g., Hawthorne et al., 2005). However in the fresh and marine water column, mobile sorbents such as plankton and particulate organic carbon can comprise the dominant sorbing phase (e.g., Schulz-Bull et al., 1998). Some of the above examples of mobile sorbing phases have been shown to play predominant role with respect to hydrocarbon bioavailability and biodegradation. Hydrocarbons associate with these matrices via the same set of sorption mechanisms as discussed above (Luthy et al., 1997). Therefore, their role in terms of reducing the bioavailability of hydrocarbons can be understood using the same terms of reference. However, their small size and thus high surface area to volume ratio implies that the exchange kinetics are more rapid (Poerschmann et al., 1997), and thus processes such as retarded diffusion play less of a bioavailability limiting role. Their small size also enables them to readily move with, for example, the advective flow of water. This, together with their propensity to associate with cellular membranes, confers on them their peculiar role in the bioavailability and biodegradation of hydrocarbons. Exactly what role do such mobile hydrocarbon sorbing phases have with respect to bioavailability? They can alter both the kinetics of abiotic mass transfer and the final phase distribution between the (stationary) sorbed and aqueous compartments. In many environments, bioavailability and thus biodegradation is limited by the slow hydrocarbon mass transfer from the sorbed state and into the aqueous phase (Harms and Bosma, 1997). Here, the stationary sorbing material can be anything from a geosorbent such as particulate matter to a non-aqueous phase liquid (NAPL). The presence of an additional sorbing phase in the surrounding aqueous medium can enhance the rate of dissolution. Both natural and synthetic surfactants increase the dissolution rate of hydrocarbons such as PAHs from their pure solid state (Grimberg et al., 1995) or when present in NAPLs (Garcia-Junco et al., 2001). In part this increase can be explained by sorption to the aqueous phase surfactant micelles reducing the dissolved phase concentrations, thus maintaining the high chemical activity gradients driving the dissolution process. However, it also appears that in parallel the surfactant micelles function as carriers, enhancing the transport of the hydrocarbons into the bulk solution (Grimberg et al., 1995). Although most information in this regard exists for biosurfactants, the nature of the mechanisms involved implies that these might also be relevant to other classes of mobile sorbents. Indeed, dissolved organic matter associated with minerals has been shown to enhance the dissolution of PAHs from NAPLs (Garcia-Junco et al., 2003). An enhancement in the dissolution rates has two implications for bioavailability. Firstly, the rate of mass transfer out of the non-accessible phase is increased which can be particularly important for those more hydrophobic hydrocarbons where dissolution is very low to start with. Secondly, the total amount of hydrocarbon in the aqueous phase can be increased above solubility, albeit partly sorbed. Should a fraction of this sorbed aqueous amount be accessible by the degrading microorganisms then this could have a positive overall effect on bioavailability.

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When considering natural environments, the distribution of hydrocarbons is generally heterogeneous. In soils and sediments, such micro scale inhomogeneities are particularly important in lowering the overall bioavailability. Since microbial colonies are spatially distributed and mainly exist attached to various surfaces rather than suspended in the interstitial solution, they ‘‘see’’ a relatively small volume of the environment (Postma and Vanveen, 1990). Should this volume become depleted of hydrocarbon via consumption, then it is rapidly the case that the distance to a replenishing source becomes too large for a sufficient re-supply via solely aqueous diffusion (Bosma et al., 1997). Therefore, hydrocarbon association with an advectively transported sorbing phase such as dissolved organic matter could also have implications for redistribution, functioning at the micro scale as ‘‘carriers’’ from a site of contamination to that of biodegradation. When considering the aquatic ecosystem, the more thorough mixing of the water column by turbulence probably means that such micro scale inhomogeneities are less of an issue. Nevertheless, in some cases heterogeneity might also play a role but over a larger scale. An oil spill at the water surface is an example. Oil hydrocarbon sorption to suspended mineral and organic matter in the water column initially might enhance the dissolution process, and then be transported away via the water currents to more distant locations, forming a plume of bioavailable hydrocarbon. Although the relevance of such roles remains to be demonstrated in the field, these are the general principles behind bioremediation of soils using surfactant washing solutions.

6

Contaminant Aging and Release Kinetics

Contaminants in geomaterials may undergo changes that have been summarized as processes of contaminant aging or weathering. It has been observed that the efficiency of chemical extraction and biodegradation of contaminants is lower when the contact time between contaminant and the geomaterials before these interventions was longer. In many cases, recent contaminations may thus be treated with higher efficiency than historical ones (Hatzinger and Alexander, 1995). For instance it has been frequently seen that when the biodegradation of a historical contamination in soil has come to an end at a still relatively high residual concentration, contaminant of the same kind that is freshly spiked to the same soil is rapidly degraded (Valo and Salkinoja-Salonen, 1986). From this experiment and similar real-world observations it can be inferred that the different portions of the contaminant show different degrees of bioaccessibility. This differential behavior of contaminant fractions is not only a problem for the remediation process but also limits the value of spiking soil with radioactively labeled contaminant as an indicator of the biodegradation potential. Which kinds of mechanisms can lead to reduced contaminant bioaccessibility and extractability? Categories of mechanisms are chemical changes, changes of the soil or sediment structure and shifts in the spatial distribution of the chemical due to diffusive transfer in combination with the exclusion of microorganisms (or extractants) from certain parts of the geomaterial. An example for chemical changes would be the successive occupation of high-energy sorption sites by contaminant molecules. The probability that individual contaminant molecules that are initially absorbed in ‘‘soft’’ organic matter of NAPL come into contact with either ‘‘hard’’ organic matter, high-energy sorption sited on the surface of e.g., soot or enter the swollen interlayers of clay minerals increases with time. Equal consequences for declining reversibility of sorption would also arise from the metamorphosis of organic matter into forms that retain absorbed molecules more efficiently. Structural changes in

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the soil matrix could lead to contaminant aging by burying formerly labile contaminant pools under poorly permeable phases. The encapsulation of ‘‘soft’’ organic matter by ‘‘hard’’ organic matter exemplified by Luthy et al. (1997) or by mineral soil constituents would be examples. One can easily imagine that bio-turbation or processing of soil materials inside of the digestive tracts of soil dwelling organisms could lead to the obstruction of diffusion pathways. The successive formation of poorly permeable interphases around tar globules would also fall into the category of structural changes. But aging can occur even in the absence of chemical or structural changes due to diffusive mass transfer. Soil and sediment contaminants typically enter via the larger pores. From thereon diffusive processes carry part of the contaminant in regions and size classes of pores that are increasingly difficult to access by microbes. Before an equilibrium distribution of the contaminant is achieved, there will be an inward-bound diffusive flux of contaminant into soil aggregates that carries contaminant fractions further away form the biota. Using experimental model polymers that exclude the possibility of chemical or structural changes, the effects of diffusion distances due to the pore-size exclusion of microbes was shown (Harms and Zehnder, 1995). The observable effect of all types of contaminant aging is an apparent desorption hysteresis, i.e., much longer time scales are needed to complete desorption than adsorption. Biodegradation curves as well as release curves obtained by continuous mild extraction (e.g., flushing with water) show pronounced tailing that may be interpreted as successive emptying of sorption sites of increasing sorption energy or as growing diffusion distances of contaminant molecules within a matrix compose of particles that empty from the outside to the center also referred to as ‘‘shrinking core’’ desorption. Mathematically, the observed desorption progress can be most easily fitted by applying first order twocompartment models that distinguish a rapidly and a slowly desorbing contaminant fraction with largely different release rate constants (Cornelissen et al., 1998). Further distinction of a third, a very slowly desorbing fraction, may give even better fits. However, seen the wealth of possible mechanisms that may bring about the earlier release of one contaminant molecule than another one, the existence of a continuum of sorption strengths and travel distances appears much more likely than the existence of two or three distinct states of sorption. The rapidly desorbing fraction has nevertheless been found to be a relatively good descriptor of the bioaccessible fraction (Cornelissen et al., 1998), whereas the extremely slowly desorbing chemically have been conceptually defined as ‘‘non-bioaccessible’’ (Semple et al., 2004) or ‘‘irreversibly-bound’’ (Reichenberg and Mayer, 2006).

7

Research Needs

In previous research, the influence of mobile sorbents and their potential to increase hydrocarbon mobility at the microscale has been largely neglected. The common opinion, that sorption generally reduces bioavailability and risk is a simplification requiring knowledgebased specification. Another blind spot of past research is the interaction of components in complex contaminants. For instance, biologically inactive components may well influence the partition behavior of bioactive components. Chemical activity-based approaches to the dynamics and fate of contaminants may thus in many cases be more appropriate than common water-solubility and concentration-based approaches.

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References Banat IM (1995) Biosurfactants production and possible uses in microbial enhanced oil-recovery and oil pollution remediation—a review. Biores Technol 51: 1–12. Bosma TNP, Middeldorp PJM, Schraa G, Zehnder AJB (1997) Mass transfer limitation of biotransformation: quantifying bioavailability. Environ Sci Technol 31: 248–252. Cornelissen G, Rigterink H, Ferdinandy MMA, van Noort PCM (1998) Rapidly desorbing fractions of PAHs in contaminated sediments as a predictor of the extent of bioremediation. Environ Sci Technol 32: 966–970. Efroymonson RA, Alexander M (1995) Reduced mineralization of low concentrations of phenanthrene because of sequestering in nonaqueous-phase liquids. Environ Sci Technol 29: 515–521. Garcia-Junco M, De Olmeda E, Ortega-Calvo JJ (2001) Bioavailability of solid and non-aqueous phase liquid (NAPL)-dissolved phenanthrene to the biosurfactant-producing bacterium Pseudomonas aeruginosa 19SJ. Environ Microbiol 3: 561–569. Garcia-Junco M, Gomez-Lahoz C, Niqui-Arroyo JL, Ortega-Calvo JJ (2003) Biosurfactant- and biodegradation-enhanced partitioning of polycyclic aromatic hydrocarbons from nonaqueous-phase liquids. Environ Sci Technol 37: 2988–2996. Ghoshal S, Luthy RG (1996) Bioavailability of hydrophobic organic compounds from nonaqueous phase liquids: the biodegradation of naphthalene from coal tar. Environ Toxicol Chem 15: 1894–1900. Grimberg SJ, Nagel J, Aitken MD (1995) Kinetics of phenanthrene dissolution into water in the presence of nonionic surfactants. Environ Sci Technol 29: 1480–1487. Harms H, Bosma TNP (1997) Mass transfer limitation of microbial growth and pollutant degradation. J Ind Microbiol Biotechnol 18: 97–105. Harms H, Zehnder AJB (1995) Bioavailability of sorbed 3-chlorodibenzofuran. Appl Environ Microbiol 61: 27–33. Hatzinger P, Alexander M (1995) Effect of aging of chemicals in soil on their biodegradability and extractability. Environ Sci Technol 29: 537–545. Hawthorne SB, Grabanski CB, Miller DJ, Kreitinger JP (2005) Solid-phase microextraction measurement of parent and alkyl polycyclic aromatic hydrocarbons in milliliter sediment pore water samples and determination of K-DOC values. Environ Sci Technol 39: 2795–2803.

Head IM, Jones DM, Ro¨ling WFM (2006) Marine microorganisms make a meal of oil. Nature Rev Microbiol 4: 173–182. Luthy RG, Aiken GR, Brusseau ML, Cunningham SD, Geschwend PM, Pignatello JJ, Reinhard M, Traina SJ, Weber WJ, Westall JC (1997) Sequestration of hydrophobic organic contaminants by geosorbents. Environ Sci Technol 31: 3341–3347. Luthy RG, Ramaswami A, Ghoshal S, Merkel W (1993) Interfacial films in coal tar nonaqueous-phase liquidwater systems. Environ Sci Technol 27: 2914–2918. Marshall AG, Rodgers RP (2004) Petroleomics: the next grand challenge for chemical analysis. Acc Chem Res 37: 53–59. McInerney MJ, Voordouw GE, Jenneman GE, Sublette KL (2007) Oil field microbiology. In Manual of Environmental Microbiology. CJ Hurst, R Crawford, JL Garland, DA Lipson, AL Mills, LD Stetzenbach (eds.). Washington, DC: ASM Press, pp. 898–911. Mozes N, Handley PS, Busscher HJ, Rouxhet PG (1991) Microbial Cell Surface Analysis: Structural and Physicochemical Methods. Verlag Chemie, Wiley, New York: Weinheim, Cambridge. Nelson EC, Ghoshal S, Edwards JC, Marsh GX, Luthy RG (1996) Chemical characterization of coal tarwater interfacial films. Environ Sci Technol 30: 1014–1022. Ortiz E, Kraatz M, Luthy RG (1999) Organic phase resistance to dissolution of polycyclic aromatic hydrocarbon compounds. Environ Sci Technol 33: 235–242. Poerschmann J, Zhang ZY, Kopinke FD, Pawliszyn J (1997) Solid phase microextraction for determining the distribution of chemicals in aqueous matrices. Anal Chem 69: 597–600. Postma J, Vanveen JA (1990) Habitable pore-space and survival of Rhizobium-Leguminosarum Biovar Trifolii introduced into soil. Microb Ecol 19: 149–161. Ramaswami A, Ghoshal S, Luthy RG (1997) Mass transfer and bioavailability of PAH compounds in coal tar NAPL-slurry systems. 2. Experimental evaluations. Environ Sci Technol 31: 2268–2276. Reichenberg F, Mayer P (2006) Two complementary sides of bioavailability: accessibility and chemical activity of organic contaminants in sediments and soils. Environ Toxicol Chem 25: 1239–1245. Schluep M, Ga¨lli R, Imboden DM, Zeyer J (2002) Dynamic equilibrium dissolution of complex nonaqueous phase liquid mixtures into the aqueous phase. Environ Toxicol Chem 21: 1350–1358. Schulz-Bull DE, Petrick G, Bruhn R, Duinker JC (1998) Chlorobiphenyls (PCB) and PAHs in water masses

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of the northern North Atlantic. Marine Chem 61: 101–114. Schwarzenbach RP, Gschwend PM, Imboden DM (2003) Environmental Organic Chemistry, 1st edn. Hoboken, NJ: Wiley. Semple K, Doick K, Jones K, Burauel P, Craven A, Harms H (2004) Defining bioavailability and bioaccessibility of contaminated soil and sediment is complicated. Environ Sci Technol 15: 229A–231A.

Valo R, Salkinoja-Salonen M (1986) Bioreclamation of chlorophenol-contaminated soil by composting. Appl Microbiol Biotechnol 25: 68–75. Volkering F, Breure AM, Sterkenburg A, van Andel JG (1992) Microbial degradation of polycyclic aromatic hydrocarbons: effect of substrate availability on bacterial growth kinetics. Appl Microbiol Biotechnol 36: 548–552.

45 Microorganism-Hydrophobic Compound Interactions H. Harms1,* . K. E. C. Smith2 . L. Y. Wick1 1 Department of Environmental Microbiology, UFZ, Helmholtz Centre for Environmental Research, Leipzig, Germany *[email protected] 2 Department of Environmental Chemistry and Microbiology, The National Environmental Research Institute (NERI), Roskilde, Denmark 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1480 2 Adhesion to Hydrocarbons as Microbial Adaptive Response . . . . . . . . . . . . . . . . . . . . . . . 1481 3 Forces and Microbial Characteristics Mediating Attachment to Hydrocarbons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1483 4 Accumulation of Hydrophobic Compounds in Bacterial Membranes and Cell-Walls . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1483 5 Is Dissolution of Hydrophobic Compounds in Water Required for Microbial Uptake? . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1484 6 Bacterial Interactions with Hydrocarbons in Mobile Sorbents . . . . . . . . . . . . . . . . . . . . 1485 7 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1488

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_101, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: The low solubility of hydrocarbons forces hydrocarbon-degrading microorganisms to physically interact with oil phases. This has various implications for the physicochemical characteristics of these microbes, their modes of hydrocarbon uptake, and behavioral and physiological strategies used to establish the interaction as summarized in > Table 1.

1

Introduction

There is consensus in the literature that water-dissolved chemicals are available to microbes (Harms and Wick, 2004). Hydrocarbons in the environment, however, occur predominantly as pure water-immiscible liquids or solids, dissolved in nonaqueous liquids, volatilized in gases, absorbed in solids, or bound to interfaces (adsorbed), whereas only tiny, often negligible fractions are dissolved in the aqueous phase. The question thus arising is ‘‘to what degree are hydrocarbons in the dominant, not water-dissolved, forms bioavailable for microbial transformation?’’ One can turn the question around and ask whether mono-disperse dissolution of hydrophobic compounds in water is required for microbial uptake or whether direct contact with substrate in other forms could allow for additional uptake. In this context, it is important

. Table 1 Overview of the different microbial strategies employed to enhance the bioavailability of nonaqueous phase liquid (NAPL) hydrocarbons to micro-organisms Category

Strategy

Consequence

Reference

Passive

Degradation in the High gradient in chemical activity maintained Harms and bulk aqueous phase Zehnder (1994)

Positional

Adhesion to interface/biofilm formation

Interaction with the NAPL

Positioning in region of highest chemical activity and reduced diffusion distance

Wick et al. (2001)

Additional diffusive barrier

Wick et al. (2001)

Chemotaxis

Positioning in region of highest chemical activity and reduced diffusion distance

Marx and Aitken (2000)

Surfactant production

Break-up of NAPL phase increasing surface area for mass transfer and colonization

Barkay et al. (1999)

Increased micellar concentrations in the aqueous phase

Volkering et al. (1995)

Direct bioavailability of micellar hydrocarbons Guha and Jaffe´ (1996a) Change in bacterial surface hydrophobicity leading to altered attachment to interfaces

Wick et al. (2002)

Inhibition of cell attachment

Neu (1996)

The literature references are those studies where the strategies have been shown to alter the hydrocarbon bioavailability

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45

to realize that bacteria:hydrophobic compound interactions take place at an extended interphase rather than at a sharp interface (Ko¨ster and van Leeuwen, 2004). The concept of an interphase considers that microorganisms are surrounded by so-called boundary layers characterized by a gradual transition from the biological to the environmental phase. From a microbe’s perspective, the actual process of uptake of a chemical typically includes transport in the aqueous medium (i.e., the boundary layer around the bacterial cell), adsorption to and absorption into extracellular hydrogel-like cell wall components, and uptake by transfer through the cell membrane, which may be followed by bioconversion or (nonconsumptive) bioaccumulation (Ko¨ster and van Leeuwen, 2004). The availability of a chemical for degradation is thus controlled by the rate of mass transfer to microbial cells relative to their intrinsic catabolic activity (Bosma et al., 1997). As the chemical transport occurs by concentrationgradient driven diffusion, relative locations of source and sink are of importance. This means that regardless of the uptake mechanism, bacterial attachment to solid, liquid, or sorbed substrates is already a powerful way to maximize substrate mass transfer. According to Fick’s law, J ¼ Deff  A @c=@x, the diffusive mass flux of a substrate toward the cell surface J is strongly affected by the space coordinate in direction of the transport @x, the area A through which the compound is diffusing, and @c, the concentration difference between the location of bioconversion and the substrate source (> Fig. 1a). The reduction of the distance between a substrate and the microorganism thus enhances diffusive mass transfer thereby increasing the availability of the hydrocarbon. For solid and liquid non aqueous phase compounds, the so-called boundary layer diffusion is the limiting step in the dissolution (Bird et al., 1960) thus controlling the rate of appearance of molecules in the bulk water phase. Even under conditions of percolation or mild stirring, particles (and bacteria) are surrounded by a diffusive boundary layer of the thickness 10–5–10–4 m (Levich, 1962). A bulk water phase containing a subequilibrium concentration of the solute will thus drive desorption from this distance resulting in a concentration gradient between the bulk phase concentration and the interfacial aqueous pseudo-equilibrium concentration. When attached to a solid, dissolved, liquid, or sorbed hydrocarbon, bacteria drive desorption by substrate uptake from inside of the boundary layer, which may result in an up to roughly hundred-fold steeper concentration gradient and mass transfer to the cell (> Fig. 1b). It has thus been concluded that the observation of lower desorption rates determined in the absence of bacteria compared with degradation rates can lead to wrong interpretations regarding the mechanisms of uptake, unless distance effects (the possibility that bacteria invade diffusion boundary layers) are accounted for (Harms and Zehnder, 1995).

2

Adhesion to Hydrocarbons as Microbial Adaptive Response

Seeing its potential effects, it is thus not surprising that bacterial adhesion to and the formation of biofilms on hydrocarbon phases have been frequently reported for transferlimited conditions in the presence of solid (e.g., Mulder et al., 1998) and liquid hydrocarbons (e.g., Rosenberg and Rosenberg, 1981). Further evidence for the importance of the close contact to the substrate comes from the observed drastic loss of microbial activity when attachment to the substrate source was suppressed with nontoxic surfactants or when adhesion-hindered mutants were used (e.g., Efroymson and Alexander, 1991; Rosenberg and Rosenberg, 1981). Bacteria also may adapt to the presence of hydrophobic (or from their perspective lipophilic) hydrocarbons by changing their membrane and outer cell wall

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. Figure 1 Conceptualized outline of the physically relevant layers at the microorganism/medium (a) and the microorganism/hydrocarbon interphase (b). Diffusive hydrocarbon flux towards the cell surface is strongly affected by the space coordinate in direction of the transport @x between the cell’s location of bioconversion of and the substrate source. When attached to a solid, dissolved, liquid or sorbed hydrocarbon, bacteria drive desorption by substrate uptake from inside of the boundary layer, which may result in an up to roughly hundred-fold steeper concentration gradient and mass transfer to the cell (b).

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properties, e.g., in order to better attach to hydrophobic surfaces (Johnsen et al., 2005; van Loosdrecht et al., 1990).

3

Forces and Microbial Characteristics Mediating Attachment to Hydrocarbons

The adhesion of bacteria to interfaces is believed to be mediated by long-ranging colloidal interactions, which attract bacteria and hold them at a close proximity of the interface, thereby facilitating short-ranging hydrophobic and electrostatic interactions to become effective and enabling the subsequent adsorption of extracellular polymeric substances, e.g., mediated by hydrogen bonds (Jucker et al., 1997; van Loosdrecht et al., 1990). It is important to note that electrostatic forces will be shielded by high ion concentrations in sea water, whereas they may play an important role in groundwater (Busscher et al., 1995). In the case of adhesion to liquids, which do not expose a rigid surface, the interaction forces will induce changes in the interface topography to optimize the interaction. It is thus to be expected that more hydrophobic bacteria will establish a larger interfacial area with a liquid hydrocarbon. Besides the establishment of more interfacial area for substrate transfer, the effect of separation distance on mass transfer (already discussed) may explain higher hydrophobicity of organisms degrading hydrophobic compounds. It has been observed, for instance, that more hydrophobic bacteria were enriched when hydrophobic membranes were used to isolate degraders of polycyclic aromatic hydrocarbons (PAH) from soil compared with conventional water extraction protocols (Bastiaens et al., 2000). The physiological role of the hydrophobicity of these organisms was supported by the observation that the PAH-degrading mycobacteria modulated their surface hydrophobicity depending on the hydrophobicity of their growth substrate by varying the length of their mycolic acids (Wick et al., 2002). It should be noted, however, that hydrophobicity is not the only cell surface characteristic controlling the contact with hydrocarbon phases. Hydrocarbon–water interfaces tend to be strongly negatively charged, which makes bacterial charge a further important determinant of the interaction. The origin of the charge is not entirely clear, but preferred adsorption of OH as opposed to H3O+ at hydrophobic liquids and solids (e.g., Teflon and polystyrene surfaces) has been suggested as the cause.

4

Accumulation of Hydrophobic Compounds in Bacterial Membranes and Cell-Walls

Various physiological adaptations of bacteria to the lipophilicity of hydrocarbons and the concomitant potential toxicity have been described (for an excellent review see Sikkema et al., 1995). Adaptations include (1) changes in the membrane fluidity by modifications of the membrane fatty acid composition, (2) changes in the membrane’s protein content and composition, (3) active excretion of toxic hydrocarbons by energy-consuming transport systems (e.g., multidrug resistance system), (4) increase of the cross-linking between the cell-wall constituents and changes in the cell wall hydrophobicity and charge, (5) modifications of the lipopolysaccharides (LPS) of the outer membrane of gram negative bacteria, and/ or (6) biotransformation and of the compound. Changes in the cell wall properties have also been described to increase the accumulation of hydrophobic compounds (e.g., Klein et al., 2008).

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Sikkema et al. (1994), correlated the n-octanol-water partition constants KOW of hydrophobic organic contaminants to their membrane-buffer partitioning coefficient KB as log KB ¼ 0:97 logKOW  0:64). This underlines the fugacity-driven bioaccumulation of hydrophobic substances in cell membranes, cell walls, and the cytoplasm. Bioaccumulation of lipophilic compounds in the lipid bilayer as a result may thus enhance their availability for biotransformation and toxic response. Accumulation in biofilms as possible sink for hydrocarbon solutes has also been described. Despite the hydrogel-like character (96–98% of water), extracellular polymeric substances of biofilms were found to contain about 60–70% of the total monoaromatic hydrocarbons isolated from the total biomass (Spa¨th et al., 1998). Bioaccumulation of hydrophobic compounds appears to be a passive process, and thermodynamically speaking, is driven by the high fugacity of hydrophobic compounds in the waterphase that promotes their expulsion from water. It is thus likely that hydrophobic cell envelopes act as transient reservoirs (biosolvents) of hydrophobic substances. Despite the normal passive character of hydrocarbon uptake, different microbial adaptations have evolved to augment uptake rates of hydrophobic organic compounds (HOC): For instance, it has been postulated that outer-membrane related lipopolysaccharides are released to encapsulate hydrocarbon droplets and to increase the efficiency of alkane mass transfer (Witholt et al., 1990). Lipopolysaccharides may influence the mass transfer via one or several of a large number of possible mechanisms, including the improvement of the dissolution process, dispersion of the soil matrix, dispersion of nonaqueous liquid phase and crystalline contaminants (Neu, 1996), the solubilization in micelles that may act as kinetic contaminant carriers through the diffuse boundary layer (Garcia et al., 2001), or thermodynamically as an additional disperse phase, or ‘‘biosolvent’’ (Noordman et al., 1998).

5

Is Dissolution of Hydrophobic Compounds in Water Required for Microbial Uptake?

This brings us to the question whether mono-disperse dissolution of hydrophobic compounds in water is a prerequisite for microbial uptake. Although it is generally accepted that substrates that occur in pure or matrix-sorbed form have to dissolve or to desorb, in order to become available for microbial uptake (for review see van Loosdrecht et al., 1990), over the past 20 years several observations have been interpreted as evidence for endocytotic uptake of ‘‘undissolved’’ substrate; this is, however, experimentally difficult to verify (Taylor and Simkiss, 2004). Three microbial uptake mechanisms of nonaqueous phase liquids have been postulated: First, entire droplets of these liquids may be channeled through pores in the cell envelope into the cytoplasm. Second, entire droplets could enter the cell surrounded by parts of the cell membrane or the cell envelope (pinocytosis) from where it can be slowly released into the cytoplasm or be degraded by membrane-bound enzymes (Taylor and Simkiss, 2004). Third, it is imaginable that droplets may fuse with and flow into the lipid bilayer upon contacting a cell membrane. Microbial uptake of solid or sorbed substrates is somewhat different. Similar to the process of pinocytosis solid substrates could directly be taken up by phagocytosis (Taylor and Simkiss, 2004). To our knowledge, however, this has never been observed and seems unlikely for bacteria, which are covered by rigid cell walls. On the other hand, solid-state molecules might directly dissolve in the membranes of contiguous bacteria upon physical contact excluding the water phase (e.g., Southam et al., 2001). The conceptual difference between adsorbed molecules and solid substrates is that the latter are

Microorganism-Hydrophobic Compound Interactions

45

adsorbed to molecules of their own kind, and with respect to their bioavailability and possibility of uptake may behave like a solid substrate. Bio-physically speaking the problem reduces to the questions whether bacteria are able (1) to enzymatically attack molecules that are still sorbed while the enzyme-substrate-complex is formed, (2) to take up molecules that are still sorbed while the transporter-substrate-complex is formed, and/or (3) to passively take up molecules without a desorption step into the aqueous phase involved (Harms and Wick, 2004). In case of absorbed molecules, pore-sequestered molecules have to move to the sorbent–water interface and desorb (i.e., dissolve) into the water phase, before film diffusion in the bulk water, and subsequent microbial uptake can take place. Intrasorbent transfer to the sorbent–water interface has often been found to control the overall desorption rate of absorbed or pore-sequestered compound. Direct contact of bacteria with absorbed hydrocarbons is thus unlikely before they appear at the sorbent–water interface and uptake of molecules by microorganisms in others than the water-dissolved state may be the exception rather than the rule (for a review cf. (Harms and Wick, 2004)). At present, there seems to be insufficient evidence for the direct uptake of hydrocarbon droplets by prokaryotes. More evidence appears to exist for the direct uptake of hydrocarbons from mobile sorbents, which will be addressed in the following section.

6

Bacterial Interactions with Hydrocarbons in Mobile Sorbents

The interaction of cells with hydrocarbons present in mobile sorbents has been rarely addressed (surfactants being an exception) and is therefore given specific attention in this section. With respect to hydrocarbon bioavailability, such mobile sorbent phases (surfactant or dissolved organic carbon; DOC) possess the following relevant characteristics: (1) propensity to sorb hydrocarbons, (2) small size and large surface area to volume ratio, (3) mobility with, for example, the flow of water, and (4) ability to interact with biological membranes. In general, it is thought that the diffusive transfer of hydrocarbons from the dissolved pool supplies a degrading microorganism (van Loosdrecht et al., 1990). Therefore, sorption to mobile sorptive phase leads to a reduction in the dissolved concentrations and is hence expected in general to lead to a concurrent reduction in the biodegradation rates. This does not imply that this sorption removes a hydrocarbon from the biodegradable pool because of a permanent association. Rather, the reduction in the dissolved concentrations driving the diffusive uptake lowers the uptake rates so that the total biodegradation is extended over a longer period of time. Interestingly, an increasing number of studies point to other transfer pathways in addition to diffusive uptake from the dissolved phase. Under certain conditions it appears that a fraction of sorbed hydrocarbon is ‘‘directly bioavailable’’ to the degrading microorganisms, i.e., it is transferred directly from the sorbed state to the degrading cell without having to first desorb into the bulk medium followed by diffusive uptake. Primarily, evidence for this has come from studies looking at the degradation kinetics of PAHs in the presence of artificial and natural surfactants (Brown, 2007; Guha and Jaffe´, 1996a, b; Guha et al., 1998). The role of surfactants in the biodegradation of hydrocarbons has been vigorously studied and found to be affected by many factors, including surfactant toxicity or preferential degradation, enhanced hydrocarbon solubilization or emulsification, and altered cell surface characteristics leading to changes in attachment behavior. Therefore, it is no surprise that the results from studies using different surfactants appear at first to be contradictory, showing an

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enhancement, inhibition, or no effect on biodegradation. Nevertheless, a series of elegant studies using various nontoxic surfactants have shown that a fraction of the micellar associated PAHs is directly bioavailable to degrading bacterial cells, without first requiring to desorb into the dissolved phase (Guha and Jaffe´, 1996a; Guha et al., 1998). This led to the development and refinement of a mechanistic model detailing this micellar transport pathway (Brown, 2007; Guha and Jaffe´, 1996b). Here, it is envisaged that the PAHs associated with the micelles are transported to, and subsequently fuse with, a hemicellar surfactant layer covering the bacterial surface, thus directly releasing the intercalated PAHs to the cell. This is shown schematically in > Fig 2a. Of course such a micelle-mediated direct PAH delivery pathway depends on the micelle forming properties of surfactants. Nevertheless, studies with other types of sorbents that are also mobile show enhanced PAH degradation kinetics, pointing to the existence of additional transfer pathways to diffusive uptake from the dissolved phase. Barkay et al. (1999) investigated the

. Figure 2 Suggested mechanisms by which a mobile sorbent phase might enhance the mass transfer of a hydrocarbon to degrading bacteria. The mechanism described in (a) is based on that described in Guha and Jaffe´, 1996b.

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45

effect of the bioemulsifier alasan on the biodegradation kinetics of phenanthrene and fluoranthene. Mineralization rates of both PAHs were enhanced by the presence of increasing amounts of alasan, indicating an enhanced mass flux to the degrading cells. Alasan is a tightly bound complex of anionic polysaccharides and proteins and forms multimolecular aggregates rather than micelles. Various studies with humic acids (HA) also point to a possible carrier role of these sorbents. Ortega-Calvo and Saiz-Jimenez (1998) looked at the mineralization of phenanthrene in batch cultures, and found that high HA loadings increased the mineralization rates, despite a reduction in the dissolved aqueous concentrations due to sorption. Laor et al. (1999) looked at the effect of sorption to Aldrich HA on the biodegradation of phenanthrene. A reduction in dissolved concentrations with increasing amounts of HA did not result in appreciable differences in the rate or extent of mineralization. This would suggest that an additional mass transport pathway offset the lowered diffusive transfer from the dissolved phase. Holman et al. (2002) studied in situ the influence of Elliott Soil HA on the degradation of a thin film of pyrene deposited on a magnetite surface. HA shortened the onset of biodegradation. These studies indicate that the association of a hydrocarbon to certain types of organic matter can, in some cases, even lead to an enhancement in the biodegradation rates. These sorbents all have the common property that they are mobile and/or can intimately associate with the microorganism surface. This appears to lead to an efficient transfer of hydrocarbon from the sorbed state to the cell. However, the nature of the mechanisms behind these observations is not always clear, despite this being a widespread phenomenon (different types of sorbing phases, different microorganisms etc). Indeed, a very different example is given by the mass transfer of PAHs between silicone polymers across an aqueous boundary layer (Mayer et al., 2005). This was enhanced in the presence of humic acids well above that resulting from molecular diffusion through the aqueous phase. The very lack of a unifying framework to explain these observations also makes this a good point in time to speculate about the mechanisms that might explain such an enhanced flux. In the case of surfactants, the fusion of hydrocarbon-filled micelles with a cell surface hemimicellar layer and direct transfer (> Fig. 2a) is well backed up by experimental evidence (Brown, 2007; Guha and Jaffe´, 1996b). However, such a mechanism is also plausible for hydrocarbons sorbed to dissolved organic matter. Dissolved organic matter is composed of low molecular weight components, which form dynamic associations via hydrophobic interactions and hydrogen bonds (Buffle and Leppard, 1995; Sutton and Sposito, 2005). Furthermore, dissolved organic matter associates with living membranes (Maurice et al., 2004), under certain conditions even leading to changes in the membrane permeability (Vigneault et al., 2000). Therefore, for a transfer pathway analogous to that of surfactants to operate, the necessary prerequisites exist. These include the surfactant-like aggregation of smaller units and adsorption to the biological membrane. This, however, has not been experimentally verified. The fact that dissolved organic matter can intimately associate with biological membranes indicates that close contact is indeed possible. In the case of silicone polymers, the direct contact between a PAH loaded donor and empty receiver compartment resulted in a very effective transfer, exceeding the transfer across an aqueous boundary layer by several orders of magnitude (Meyer et al., 2005, 2007). Therefore, it is feasible that should the collision between bacteria and dissolved organic matter (DOM) lead to direct contact, albeit temporary, this could also lead to the transfer of hydrocarbon from sorbent to cell (> Fig. 2b). For many hydrocarbons, the main resistance to diffusive transfer to/from an organic-type matrix lies in the aqueous boundary layer (Schwarzenbach et al., 2003). This implies that the chemical

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activity in this layer is below that in the bulk solution. Therefore, if a mobile sorbent phase, at equilibrium with the bulk solution, enters this depleted boundary layer, desorption could occur. The extent would depend on the residence time in this layer and the kinetics of hydrocarbon release (> Fig. 2c).

7

Research Needs

One challenge is to identify the nature of the mechanisms behind the often observed increase in direct bioavailability to degrading microorganisms of carrier-sorbed hydrocarbons. This knowledge would contribute to understanding those situations where direct hydrocarbon transfer plays a dominating role in bioavailability. The application of the newest generation of powerful microscopic techniques such as two-photon microscopy or high-resolution light microscopy may offer ways to obtain more mechanistic insight into uptake processes. Finally it is still unclear exactly what role the sorption to other mobile phases such as bacteria has. In a bacterial bloom developing after a spill of crude oil, the dominating bacterial biomass may not be involved in the biodegradation of specific hydrocarbons. (Head et al., 2006). Nevertheless, this may function as a large sorptive phase, but any role in bioavailability is unclear.

References Barkay T, Navon-Venezia S, Ron EZ, Rosenberg E (1999) Enhancement of solubilization and biodegradation of polyaromatic hydrocarbons by the bioemulsifier alasan. Appl Environ Microbiol 65: 2967–2702. Bastiaens L et al. (2000) Isolation of adherent polycyclic aromatic hydrocarbon (PAH) degrading bacteria using PAH sorbing carriers. Appl Environ Microbiol 66: 1834–1843. Bird RB, Stewart WE, Lightfoot EN (1960) Transport Phenomena,1st edn. New York: Wiley. Bosma TNP, Middeldorp PJM, Schraa G, Zehnder AJB (1997) Mass transfer limitation of biotransformation: Quantifying bioavailability. Environ Sci Technol 31: 248–252. Brown DG (2007) Relationship between micellar and hemi-micellar processes and the bioavailability of surfactant-solubilized hydrophobic organic compounds. Environ Sci Technol 41: 1194–1199. Buffle J, Leppard GG (1995) Characterization of aquatic colloids and macromolecules. 1. Structure and behavior of colloidal material. Environ Sci Technol 29: 2169–2175. Busscher HJ, van de Beltgritter B, van der Mei HC (1995) Implications of microbial adhesion to hydrocarbons for evaluating cell-surface hydrophobicity: 1. Zeta potentials of hydrocarbon droplets. Colloids Surf B Biointerfaces 5: 111–116.

Efroymson RA, Alexander M (1991) Biodegradation by an Arthrobacter species of hydrocarbon partitioned into an organic solvent. Appl Environ Microbiol 57: 1441–1447. Garcia JM, Wick LY, Harms H (2001) Influence of the nonionic Surfactant Brij 35 on the bioavailability of solid and sorbed dibenzofuran. Environ Sci Technol 35: 2033–2039. Guha S, Jaffe´ PR (1996a) Biodegradation kinetics of phenanthrene partitioned into the micellar phase of nonionic surfactants. Environ Sci Technol 30: 605–611. Guha S, Jaffe´ PR (1996b) Bioavailability of hydrophobic compounds partitioned into the micellar phase of nonionic surfactants. Environ Sci Technol 30: 1382–1391. Guha S, Jaffe´ PR, Peters CA (1998) Bioavailability of mixtures of PAHs partitioned into the micellar phase of a nonionic surfactant. Environ Sci Technol 32: 2317–2324. Harms H, Wick LY (2004) Mobilization of organic compounds and iron by microorganisms. In Physicochemical Kinetics and Transport at Chemical– Biological Interphases. HP van Leuven, W Koester (eds.). Chichester: Wiley, pp. 401–444. Harms H, Zehnder AJB (1994) Influence of substrate diffusion on degradation of dibenzofuran and

Microorganism-Hydrophobic Compound Interactions 3-chlorodibenzofuran by attached and suspended bacteria. Appl Environ Microbiol 60: 2736–2745. Harms H, Zehnder AJB (1995) Bioavailability of sorbed 3-chlorodibenzofuran. Appl Environ Microbiol 61: 27–33. Head IM, Jones DM, Roling WFM (2006) Marine microorganisms make a meal of oil. Nature Rev Microbiol 4: 173–182. Holman HN, Nieman K, Sorensen DL, Miller CD, Martin MC, Borch T, McKinney WR, Sims RC (2002) Catalysis of PAH biodegradation by humic acid shown in synchrotron infrared studies. Environ Sci Technol 36: 1276–1280. Johnsen AR, Wick LY, Harms H (2005) Principles of microbial PAH degradation. Environ Poll 133: 71–84. Jucker BA, Harms H, Hug SJ, Zehnder AJB (1997) Adsorption of bacterial surface polymers on mineral oxides is mediated by the formation of hydrogen bonds. Colloids Surf B Biointerfaces 9: 331–343. Klein B, Grossi V, Bouriat P, Goulas P, Grimaud R (2008) Cytoplasmic wax ester accumulation during biofilm-driven substrate assimilation at the alkane – water interface by Marinobacter hydrocarbonoclasticus SP17. Res Microbiol 159: 137–144. Ko¨ster W, van Leeuwen HP (2004) Physicochemical kinetics and transport at the biointerface: setting the stage. In Physicochemical kinetics and transport at biointerfaces. W Ko¨ster, HP van Leeuven (eds.). Chichester (GB): Wiley. Laor Y, Strom PF, Farmer WJ (1999) Bioavailability of phenanthrene sorbed to mineral-associated humic acid. Water Res 33: 1719–1729. Levich V (1962) Physicochemical Hydrodynamics. Englewood, NJ: Prentice-Hall. Luthy RG, et al. (1997) Sequestration of hydrophobic organic contaminants by geosorbents. Environ Sci Technol 31: 3341–3347. Marx RB, Aitken MD (2000) Bacterial chemotaxis enhances naphthalene degradation in a heterogenous aqueous system. Environ Sci Technol 34: 3379–3383. Maurice PA, Manecki M, Fein JB, Schaefer J (2004) Fractionation of an aquatic fulvic acid upon adsorption to the bacterium, Bacillus subtilis. Geomicrobiol J 21: 69–78. Mayer P, Fernqvist MM, Christensen PS, Karlson U, Trapp S (2007) Enhanced diffusion of polycyclic aromatic hydrocarbons in artificial and natural aqueous solutions. Environ Sci Technol 41: 6148–6155. Mayer P, Karlson U, Christensen PS, Johnsen AR, Trapp S (2005) Quantifying the effect of medium composition on the diffusive mass transfer of hydrophobic organic chemicals through unstirred boundary layers. Environ Sci Technol 39: 6123–6129.

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Mulder H, Breure AM, van Honschooten D, Grotenhuis JT, van Andel JG, Rulkens WH (1998) Effect of biofilm formation by Pseudomonas 8909N on the bioavailability of solid naphthalene. Appl Microbiol Biotechnol 50:277–283. Neu TR (1996) Significance of bacterial surface-active compounds in interaction of bacteria with interfaces. Microbiol Rev 60: 151–166. Noordman WH, Ji W, Brusseau ML, Janssen DB (1998) Effects of rhamnolipid biosurfactants on removal of phenanthrene from soil. Environ Sci Technol 32: 1806–1812. Ortega-Calvo JJ, Saiz-Jimenez C (1998) Effect of humic fractions and clay on biodegradation of phenanthrene by a Pseudomonas fluorescens strain isolated from soil. Appl Environ Microbiol 64: 3123–3126. Rosenberg M, Rosenberg E (1981) Role of adherence in growth of Acinetobacter cacoaceticus RAG-1 on hexadecane. J Bacteriol 148: 51–57. Schwarzenbach RP, Gschwend PM, Imboden DM (2003) Environmental Organic Chemistry, 1st edn. Hoboken (NJ): Wiley. Sikkema J, Debont JAM, Poolman B (1994) Interactions of cyclic hydrocarbons with biological-membranes. J Biol Chem 269: 8022–8028. Sikkema J, de Bont JAM, Poolman B (1995) Mechanism of membrane toxicity of hydrocarbons. Microbiol Rev 59: 201–222. Southam G, Whitney M, Knickerbocker C (2001) Structural characterization of the hydrocarbon degrading bacteria-oil interface: implications for bioremediation. Int Biodeter Biodegr 47: 197–201. Spa¨th R, Flemming HC, Wuertz S (1998) Sorption properties of biofilms. Wat Sci Technol 37: 207–210. Sutton R, Sposito G (2005) Molecular structure in soil humic substances: The new view. Environ Sci Technol 39: 9009–9015. Taylor MG, Simkiss K (2004) Transport of colloids and particles across biological membranes. In Physicochemical Kinetics and Transport at Chemical– Biological Interphases. HP van Leeuwen, W Koester (eds.). Chichester: Wiley, pp. 358–400. van Loosdrecht MCM, Lyklema J, Norde W, Schraa G, Zehnder AJB (1990) Influence of interfaces on microbial activity. Microb Rev 54: 75–87. Vigneault B, Percot A, Lafleur M, Campbell PGC (2000) Permeability changes in model and phytoplankton membranes in the presence of aquatic humic substances. Environ Sci Technol 3: 3907–3913. Volkering F, Breure AM. van Andel JG, Rulkens WH (1995) Influence of nonionic surfactants on bioavailability and biodegradation of polycyclic

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aromatic hydrocarbons. Appl Environ Microbiol 61: 1699–1705. Wick LY, Colangelo-Failla T, Harms H (2001) Kinetics of mass transfer-limited microbial growth on solid PAHs. Environ Sci Technol 35: 354–361. Wick LY, de Munain AR, Springael D, Harms H (2002) Responses of Mycobacterium sp. LB501T to the low

bioavailability of solid anthracene. Appl Microbiol Biotechnol 58: 378–385. Witholt B, Desmet MJ, Kingma J, van Beilen JB, Kok M, Lageveen RG, Eggink G (1990) Bioconversions of aliphatic compounds by Pseudomonas oleovorans in multiphase bioreactors – Background and economic potential. Trends Biotechnol 8: 46–52.

46 Biofilm Development at Interfaces between Hydrophobic Organic Compounds and Water R. Grimaud Institut Pluridisciplinaire de Recherche en Environnement et Mate´riaux, Equipe Environnement et Microbiologie, Universite´ de Pau et des Pays de l’Adour, Pau, France [email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1492 2 Multispecies Biofilms on Hydrophobic Interface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1494 3 Cell Adhesion to Hydrophobic Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1494 4 Regulation and Determinism of Biofilm Formation at Apolar Substrates–Water Interfaces . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1495 5 Biofilm Formation as an Adaptive Response to Optimize Acquisition of Insoluble HOCs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1496 6 Research Needs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1497

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_102, # Springer-Verlag Berlin Heidelberg, 2010

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Abstract: Hydrophobic organic compounds (HOCs) encompass a great variety of molecules, including contaminants such as hydrocarbons, which are used as substrates by bacteria (See > Chapter 20, Vol. 1, Part 4). It has been known for a long time that many bacterial strains using HOCs as carbon and energy source grow and form biofilms at the water–HOCs interface. Because they develop on an interface serving as both substratum and substrate, these biofilms represent a distinct class of biofilm exhibiting unique properties. They tend to occur specifically at the surface of nearly insoluble hydrophobic substrates, implying that nutritive interfaces exert a control on biofilm determinism and development. Furthermore, biofilms on HOCs possess the capacity to overcome the low accessibility of nearly water-insoluble substrates. These two properties make biofilm formation on HOCs a critical process in the biodegradation of hydrophobic contaminants and represent an original facet of biofilm biology.

1

Introduction

Early studies on hydrocarbon biodegradation led to the observation that hydrocarbondegrading bacteria had high affinity for oil droplets. Phase contrast and electron microscopy examination of Acinetobacter sp. growing on hexadecane revealed hydrocarbons spheres densely covered with bacterial cells and indicated direct contact between the cells and oil droplets (Kennedy et al., 1975). Since then, similar observations have been reiterated with various alkane degrading strains, Rhodococcus sp. Q15 (Whyte et al., 1999), Acinetobacter venetianus RAG-1 (Baldi et al., 1999), Oleiphilus messinensis (Golyshin et al., 2002), Pseudomonas UP-2 (Zilber Kirschner et al., 1980) and Marinobacter hydrocarbonoclasticus SP17 (> Fig. 1) (Klein et al., 2008). Attachment to polycyclic aromatic hydrocarbons (PAHs) has also been described for Pseudomonas spp (Eriksson et al., 2002; Mulder et al., 1998), Sphingomonas sp.CHY-1 (Willison, 2004) and Mycobacterium frederiksbergense LB501T (Bastiaens et al., 2000). In biphasic medium culture containing a poorly water-soluble substrate and the aqueous phase, substrate-bounds cells often coexist with cells floating freely in the aqueous phase. Although the presence of hydrocarbon bound cells at the interface assumes interfacial growth, demonstration of actual substrate degradation and growth of the attached cell has been provided in only a few cases (Efroymson and Alexander, 1991; Wick et al., 2003; Zilber Kirschner et al., 1980). The sessile mode of life and the multilayered structure of cells growing at the interface between HOCs (Hydrophobic Organic Compounds) and water are reminiscent of biofilms. During the last two decades, biofilms have been the subject of extensive investigations. Most of our knowledge of the molecular biology of biofilms has been derived from model strains such as Pseudomonas aeruginosa and Escherichia coli. This research has revealed that biofilms are much more than the simple accretion of cells attached to an interface. Biofilms are heterogeneous, highly organized structures possessing an architecture that is essential to their functioning. Biofilm growth follows a stepwise pattern of development involving cell differentiation and collective behavior of the cells (Stewart and Franklin, 2008; Webb et al., 2003). Molecular studies of bacteria growing at hydrophobic interfaces have not yet gone far enough to say whether they share all the characteristics of extensively studied model biofilms. However, two properties characteristic of the biofilm lifestyle have been identified in bacteria growing on HOCs. First, CSLM (Confocal Scanning Laser Microscopy) observation of a biofilm community developing at polychlorinated biphenyl-water interface provided evidence of a stepwise development pattern of the biofilm (Macedo et al., 2005). Simple growth at

Biofilm Development at Interfaces between Hydrophobic Organic Compounds and Water

b

a

46

50 µm

Biofilm

Eicosane

c

1 mm

. Figure 1 M. hydrocarbonoclasticus SP17 biofilms growing on alkanes. (a) and (b), confocal scanning laser microscopy images of a biofilm covered hexadecane droplet. Hydrophobic regions including bacteria as well as hydrocarbon were stained with red Nile (red signal), glycoconjugates were stained with PSA lectin (green signal). In (a), the data are presented as an isosurface projection where the two signals have been split. In (b), the dataset is presented as an XYZ projection. The two signals were not separated, colocalized signals of the green and red channel appear in yellow indicating the colocalization of Nile red stain as well as from the lectin stain. Images courtesy by Pierre-Joseph Vaysse and Thomas R. Neu (Helmholtz Centre for Environmental Research - UFZ, Magdeburg, Germany). (c), picture of M. hydrocarbonoclasticus SP17 biofilm growing at the surface of solid eicosane.

interface would not result in different stages of development. Sequential development is indicative of the execution of a genetic program and of intercellular communication. Second, Whyte et al. (1999) detected production of polysaccharide during growth at the surface of alkane droplets indicating production of extracellular matrix, which is a typical trait of biofilms. In this chapter, biofilms on HOCs refers to multilayered, matrix-embedded bacteria or bacterial communities growing at the HOCs–water interface and using these compounds as a substrate. In such biofilms, the energy and the carbon, which fuel bacterial growth, are provided by the degradation of the substrate which thus constitutes a nutritive interface serving as both substrate and substratum.

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Multispecies Biofilms on Hydrophobic Interface

During enrichment procedures on hydrocarbons, microbiologists have very often observed multispecies biofilms developing at the hydrocarbon–water interface. For example, Deppe et al., have (Deppe et al., 2005) observed by phase contrast microscopy oil droplets covered by a biofilm during isolation of a consortium enriched on crude oil from Arctic Sea ice and seawater from Spitzbergen. Unfortunately, such observations received little attention and in consequence they are not always mentioned in the literature and are rarely fully documented, making it difficult to measure the occurrence of these biofilms among bacterial species. Stach et al. (Stach and Burns, 2002) carried out a study devoted to the diversity of biofilm communities developing on PAHs. Biofilms developing on naphthalene- and phenanthrenecoated flow cells were isolated and their diversity compared with planktonic cultures enriched on the same hydrocarbons. The biofilm system produced a three times as high diversity of culturable bacteria as the enrichment culture. Molecular methods revealed that the biofilm community contained a greater diversity of actives species and of PAH degradation genes than the planktonic enrichment community. The active species diversity found in the biofilm closely matched the diversity found in the PAH-contaminated soil used as inoculum. This study demonstrates that biofilm cultures represent a mean to obtain PAH–degrading communities closely related to environmental situations suggesting that biofilm formation on hydrocarbons is a likely lifestyle in natural ecosystems.

3

Cell Adhesion to Hydrophobic Compounds

Either in order to develop a biofilm or to grow as single cell layer at the interface, bacteria must first approach and then adhere to the interface. Bacteria can reach surfaces by passive diffusion, random swimming, or taxis that is a directed motility in response to chemical and physical gradients. Chemotaxis has been observed in response to single ring aromatic hydrocarbons, naphthalene, and hexadecane. Regrettably, no experiment designed to determine whether this behavior led to biofilm (See > Chapters 49 and 50, Vol. 2, Part 8) formation at the interface between water and hydrocarbons has been conducted (Lanfranconi et al., 2003; Pandey and Jain, 2002). To date, random mobility like swimming has never been shown to play a role in adhesion to HOCs. Once cells have reached the interface, the initial adhesion step is a purely physicochemical process described by the traditional and extended DLVO theories of colloidal stability, which describes contact of cell to surface as the result of van der Waals interactions, Lewis acid–base interactions, and electrostatic interactions (Hermansson, 1999). The intensity of these interactions and hence the effectiveness of the binding depend on the cell surface properties (hydrophobicity, charge, roughness, etc.) as well as interface properties. This means that only bacteria exhibiting the proper surface properties will adhere on hydrophobic surfaces. In many cases, adhered cells exhibit surface properties that are different from their soluble substrate-grown counterparts. These alterations of the cell surface are thought to strengthen adhesion after the initial interaction with the interface. For example, anthracenegrown cells of Mycobacterium sp. LB501T are more hydrophobic and more negatively charged than glucose-grown cells (Wick et al., 2002). Changes in cell surface can be achieved by modification, production, or removal of their surface molecules. In Gram-positive bacteria, the presence and the chain length of mycolic acids are correlated with hydrophobicity and adherence (Bendinger et al., 1993). Lipopolysaccharides are important determinants of cell

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surface properties in Gram-negative bacteria. Their chain length variation or their removal from the cell surface has been shown to be important for interacting with hydrocarbons (Al-Tahhan et al., 2000). Capsular Polysaccharides are another class of surface molecule playing a role in adhesion to hydrophobic compounds (Baldi et al., 1999) (> Chapters 3, 5 and 6, Vol. 2, Part 2). Adhesion to hydrophobic surfaces is also mediated by extracellular appendages such as frimbriae and pili. The importance of fimbriae in adherence to hexadecane was demonstrated by the isolation of a nonadherent mutant of Acinetobacter calcoaceticus RAG-1 which was devoid of frimbriae and defective for growth on hydrocarbons. The reappearance of fimbriae in adherent revertants was a strong argument in favor of the involvement of fimbriae in adhesion to hexadecane (Rosenberg et al., 1982). Similarly, long, thick fimbriae have been observed on cells of Acinetobacter haemolyticus AR-46 growing at the surface of n-alkane droplets. The authors assume that these fimbriae play a major role in n-alkane utilization (Bihari et al., 2007). Changes in surface properties of cells grown on hydrophobic substrates and inhibition of adhesion in presence of soluble substrates indicate that the process of adhesion is regulated and cells are able to respond to contact with hydrophobic interfaces (Wick et al., 2002).

4

Regulation and Determinism of Biofilm Formation at Apolar Substrates–Water Interfaces

In biofilms growing on an inert substratum, all nutrients are supplied through the aqueous phase. In biofilm on HOCs, the situation is very different. These biofilms develop in a biphasic medium where the electron donor is provided by the nonaqueous phase and the electron acceptor (e.g., oxygen) is available from the aqueous phase. It results in a geometry that one side of the biofilm is in close association with the electron source, while the other contacts the source of the electron acceptor. Thus, cells within the biofilm experience two opposite gradients of acceptor and donor of electron generated by their simultaneous diffusion and consumption. These microscale chemical gradients presumably contribute to the physiological heterogeneity in the biofilm and exert a control of its development. The experiment carried out by JoannisCassan et al. (2005) demonstrated that biofilm growth on hydrocarbons can be limited either by carbon or by oxygen depletion. They studied biofilm growth in a liquid–liquid system consisting of an emulsion obtained by stirring dodecane in mineral medium. Biofilm growth occurred at the surface of a dodecane droplet. During growth, droplet diameter was reduced from 200 mm to 160 mm. Biofilm growth ceased when it reached a maximum thickness at about 80 mm. A series of experiments demonstrated that inhibition of growth was caused by the diffusion limitation of both dodecane and oxygen within the biofilm but not by others factors such as nutrients exhaustion or product inhibition (Joannis-Cassan et al., 2005). The substrate/substratum specificity of biofilm on hydrocarbons is certainly one remarkable feature that distinguishes them from other biofilms. Biofilm formation tends to occur preferentially on less soluble substrates and seems to be regulated as a function of substrate availability. Screening for biofilm formation capacity by isolated PAHs degrading strains showed that majority of the tested strains formed biofilm in microtiter wells coated with PAH crystals. For strains capable of growing on different PAHs, it was observed that the percentage of adhering cells decreases with the solubility of the PAHs, indicating that aqueous solubility of the substrate exerts a regulation on biofilm development (Johnsen and Karlson, 2004).

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Pseudomonas putida ATCC 17514 exhibits different growth patterns depending on the PAH properties on which it is feeding. CSLM observation of a gpf-labeled derivative of this strain showed that growth on phenanthrene occurred by forming a biofilm at the crystal surface, while on fluorene, which is more soluble than phenanthrene, P. putida grew randomly between the crystals feeding on the dissolved PAH (Rodrigues et al., 2005). Insoluble substrate preference for biofilm formation has also been observed in M. hydrocarbonoclasticus SP17. This bacterium forms biofilms on a variety of HOCs, including n-alkanes, wax esters, and triglycerides, but is unable to form biofilm on either nonmetabolizable compounds such as branched alkanes and n-alkane with more than 28 carbon atoms or on inert substratum such as glass or plastic in presence of acetate as substrate (Klein et al., 2008); (See > Chapter 5, Vol. 3, Part 1; > Chapter 34, Vol. 2, Part 6;) unpublished results. The preference for insoluble substrates suggests that bacteria forming biofilm on HOCs are able to detect and recognize nutritive interfaces. It is reasonable to anticipate that control of biofilm formation by substrate/substratum is exerted through a signal transduction pathway and genetic regulations. Indeed, induction of genes at an interface was demonstrated for pra gene encoding the PA protein an alkane inducible extracellular protein exhibiting an emulsifying activity involved in hexadecane assimilation and the rhlR gene coding for the transcriptional activator of rhamnolipids biosynthesis. Studies with liquid cultures on hexadecane of P. aeruginosa harboring a pra::gfp or rhlR::gfp fusion revealed specific transcriptional activity at the hexadecane–water interface (Holden et al., 2002).

5

Biofilm Formation as an Adaptive Response to Optimize Acquisition of Insoluble HOCs

The first intuitive indications that biofilms could favor access to poorly soluble HOCs came from the observations that biofilm formation is a function of substrate solubility. The more insoluble the substrate is, the more growth occurs at the water–HOC interface. Moreover, it was observed that several strains growing at the interface between nearly insoluble hydrocarbons and water do not release emulsifier or surface-active compounds in the bulk medium (Bouchez et al., 1997; Bouchez-Naı¨tali et al., 1999, 2001; Klein et al., 2008; Wick et al., 2002). In these cases, cells do not access the substrate by surfactant-mediated transfer where cells contact emulsified, solubilized or pseudo solubilized hydrocarbons. Accession to the insoluble substrate rather occurs by direct contact of the cells or extracellular structures with the hydrocarbon–water interface. Rosenberg demonstrated the importance of adhesion to hydrocarbons in the growth Acinetobacter calcoaceticus RAG-1 on hexadecane in absence of emulsifier (Rosenberg and Rosenberg, 1981). Thus, biofilm formation and adhesion to hydrocarbons would promote growth on hydrocarbons by facilitating interfacial access. The strongest evidence of increase of access to HOCs by adhesion or biofilm formation arose from kinetic studies showing that growth at the interface occurs faster than mass transfer rate of HOCs in the absence of bacteria would suggest. (Bouchez-Naı¨tali et al., 2001; Calvillo and Alexander, 1996; Harms and Zehnder, 1995; Wick et al., 2002, 2003). Mechanisms employed in biofilms for accessing HOC are still poorly understood. On the one hand, it is not difficult to imagine that biofilms offer a way to optimize the effect of known mechanisms of acquisition of poorly soluble hydrophobic substrates. Surfactant production within a biofilm would limit its dilution in the bulk phase, facilitating the formation of micelle by keeping the concentration the surfactant close to the critical micelle concentration. Biofilms also offer the advantage of holding the cell population in the vicinity of the HOC–water

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46

interface thus stimulating the mass transfer of HOC by shortening the diffusive pathway (Wick et al., 2002). On the other hand, biofilm lifestyle might offer possibilities of biofilm-specific mechanisms for HOCs accession. Biofilms are typically characterized by dense cell clusters embedded in extracellular polymeric substances. Formation of such structures involves profound changes of cell physiology and behavior requiring regulation of the expression of hundreds of genes. The metabolic activities of the cells, together with diffusion processes, generate chemical gradients within the biofilm which, in turn, affect the physiology of the cells. Consequently, biofilms are highly heterogeneous structures containing cells exhibiting various phenotypes, metabolisms, and physiological states (Stewart and Franklin, 2008). In many cases biofilm development has been shown to be controlled by intercellular signaling, or quorum sensing, indicating that cell activities are to some extent coordinated (Irie and Parsek, 2008). In view of such a reshaping of cellular function and structural organization, the existence biofilm-specific mechanisms for HOCs accession is conceivable. One example of biofilmspecific mechanism involved in HOCs assimilation is the storage of the herbicide diclofop in exopolymer matrix and its subsequent utilization by the biofilm community (Wolfaardt et al., 1995).

6

Research Needs

The most exciting aspect of biofilms on HOCs is certainly what distinguishes them from other biofilms, that is to say, their substrate/substratum specificity and their capacity to overcome the low accessibility of a hydrophobic substrate. These two properties make biofilm formation a very efficient adaptive strategy to assimilate HOCs which can provide a serious advantage in environments where carbon sources are scarce. The processes by which biofilms stimulate interfacial accession to nearly insoluble substrates remain to be elucidated. Substrate specificity of biofilm formation for nutritive interfaces presumably involves surface sensing and signal transduction pathways, which have not been revealed yet. Biofilm development during the assimilation of HOCs most likely requires coordination of fundamental processes such as architectural biofilm organization, physico-chemical interactions between biofilm and substrate, and the control of gene expression. Investigation of these processes will require multidisciplinary approaches aimed at (1) identifying the genes/proteins involved in biofilm formation, (2) deciphering the architecture of biofilms, and (3) characterizing at the physicochemical level the interactions between biofilm components (cells and extracellular matrix) with hydrophobic substrates. So far, investigations of biofilms on HOCs have been conducted on different strains growing on various substrates in diverse experimental setups. It was therefore not possible to correlate these results in order to draw a picture of the functioning of these biofilms. The use of a few model bacteria chosen for their ability to form readily and reproducibly biofilms on HOCs, their genetic amenability, and the availability of their genome sequence would ensure the complementarity of the data obtained from multidisciplinary approaches. In addition to studies examining model single species biofilms on HOCs, investigations of the activities and biodiversity of multispecies biofilms isolated from samples collected from various environments are critical to our full understanding of the ecological significance of these biofilms. Due to their wide distribution in the environment, their recalcitrance, and their deleterious effect on human health, hydrocarbons have been the main molecules targeted in studies of biofilm formation on HOCs. However, other classes of HOCs should also be taken into

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consideration. In the natural environment, lipids represent a very abundant class of HOCs. For example, in sea water they represent up to about 25% of the particulate organic carbon, the biodegradation of which is relevant to the global carbon cycle (Bendtsen et al., 2002).

References Al-Tahhan RA, Sandrin TR, Bodour AA, Maier RM (2000) Rhamnolipid-induced removal of lipopolysaccharide from Pseudomonas aeruginosa: Effect on cell surface properties and interaction with hydrophobic substrates. Appl Environ Microbiol 66: 3262–3268. Baldi F, Ivosˇevic N, Minacci A, Pepi M, Fani R, Svetlicˇic V, Zˇutic V (1999) Adhesion of Acinetobacter venetianus to diesel fuel droplets studied with in situ electrochemical and molecular probes. Appl Environ Microbiol 65: 2041–2048. Bastiaens L, Springael D, Wattiau P, Harms H, deWachter R, Verachtert H, Diels L (2000) Isolation of adherent polycyclic aromatic hydrocarbon (PAH)-degrading bacteria using PAH-sorbing carriers. Appl Environ Microbiol 66: 1834–1843. Bendinger B, Rijnaarts HHM, Altendorf K, Zehnder AJB (1993) Physicochemical cell surface and adhesive properties of coryneform bacteria related to the presence and chain length of mycolic acids. Appl Environ Microbiol 59: 3973–3977. Bendtsen J, Lundsgaard C, Middleboe M, Archer D (2002) Influence of bacterial uptake on deep-ocean dissolved organic carbon. Global Biogeochemical Cycles 16: 74–71. Bihari Z, Pettko-Szandtner A, Csanadi G, Balazs M, Bartos P, Kesseru P, Kiss I, Mecs I (2007) Isolation and characterization of a novel n-alkane-degrading strain, acinetobacter haemolyticus ar-46. Zeitschrift fur Naturforschung - Section C Journal of Biosciences 62: 285–295. Bouchez M, Blanchet D, Vandecasteele JP (1997) An interfacial uptake mechanism for the degradation of pyrene by a Rhodococcus strain. Microbiology 143: 1087–1093. Bouchez-Naı¨tali M, Blanchet D, Bardin V, Vandecasteele JP (2001) Evidence for interfacial uptake in hexadecane degradation by Rhodococcus equi: The importance of cell flocculation. Microbiology 147: 2537–2543. Bouchez-Naı¨tali M, Rakatozafy H, Leveau JY, Marchal R, Vandecasteele JP (1999) Diversity of bacterial strains degrading hexadecane in relation to the mode of substrate uptake. J Appl Microbiol 86: 421–428. Calvillo YM, Alexander M (1996) Mechanism of microbial utilization of biphenyl sorbed to polyacrylic beads. Appl Microbiol Biotechnol 45: 383–390.

Deppe U, Richnow HH, Michaelis W, Antranikian G (2005) Degradation of crude oil by an arctic microbial consortium. Extremophiles 9: 461–470. Efroymson RA, Alexander M (1991) Biodegradation by an arthrobacter species of hydrocarbons partitioned into an organic solvent. Appl Environ Microbiol 57: 1441–1447. Eriksson M, Dalhammar G, Mohn WW (2002) Bacterial growth and biofilm production on pyrene. FEMS Microbiol Ecol 40: 21–27. Golyshin PN, Chernikova TN, Abraham WR, Lunsdorf H, Timmis KN, Yakimov MM (2002) Oleiphilaceae fam. Nov., to include Oleiphilus messinensis gen. Nov., sp. Nov., a novel marine bacterium that obligately utilizes hydrocarbons. Int J Syst Evol Microbiol 52: 901–911. Harms H, Zehnder AJB (1995) Bioavailability of sorbed 3-chlorodibenzofuran. Appl Environ Microbiol 61: 27–33. Hermansson M (1999) The DLVO theory in microbial adhesion. Colloids Surf B Biointerfaces 14: 105–119. Holden PA, LaMontagne MG, Bruce AK, Miller WG, Lindow SE (2002) Assessing the role of Pseudomonas aeruginosa surface-active gene expression in hexadecane biodegradation in sand. Appl Environ Microbiol 68: 2509–2518. Irie Y, Parsek MR (2008) Quorum sensing and microbial biofilms. Curr Top Microbiol Immunol 322: 67–84. Joannis-Cassan C, Delia ML, Riba JP (2005) Limitation phenomena induced by biofilm formation during hydrocarbon biodegradation. J Chem Technol Biotechnol 80: 99–106. Johnsen AR, Karlson U (2004) Evaluation of bacterial strategies to promote the bioavailability of polycyclic aromatic hydrocarbons. Appl Microbiol Biotechnol 63: 452–459. Kennedy RS, Finnerty WR, Sudarsanan K, Young RA (1975) Microbial assimilation of hydrocarbons. I. The fine structure of a hydrocarbon oxidizing Acinetobacter sp. Arch Microbiol 102: 75–83. Klein B, Grossi V, Bouriat P, Goulas P, Grimaud R (2008) Cytoplasmic wax ester accumulation during biofilmdriven substrate assimilation at the alkane-water interface by Marinobacter hydrocarbonoclasticus SP17. Res Microbiol 159: 137–144.

Biofilm Development at Interfaces between Hydrophobic Organic Compounds and Water Lanfranconi MP, Studdert CA, Alvarez HM (2003) A strain isolated from gas oil-contaminated soil displays chemotaxis towards gas oil and hexadecane. Environ Microbiol 5: 1002–1008. Macedo AJ, Kuhlicke U, Neu TR, Timmis KN, Abraham WR (2005) Three stages of a biofilm community developing at the liquid-liquid interface between polychlorinated biphenyls and water. Appl Environ Microbiol 71: 7301–7309. Mulder H, Breure AM, Van Honschooten D, Grotenhuis JTC, Van Andel JG, Rulkens WH (1998) Effect of biofilm formation by Pseudomonas 8909n on the bioavailability of solid naphthalene. Appl Microbiol Biotechnol 50: 277–283. Pandey G, Jain RK (2002) Bacterial chemotaxis toward environmental pollutants: Role in bioremediation. Appl Environ Microbiol 68: 5789–5795. Rodrigues AC, Brito AG, Wuertz S, Melo LF (2005) Fluorene and phenanthrene uptake by Pseudomonas putida ATCC 17514: Kinetics and physiological aspects. Biotechnol and Bioeng 90: 281–289. Rosenberg M, Bayer EA, Delarea J, Rosenberg E (1982) Role of thin fimbriae in adherence and growth of Acinetobacter calcoaceticus RAG-1 on hexadecane. Appl Environ Microbiol 44: 929–937. Rosenberg M, Rosenberg E (1981) Role of adherence in growth of Acinetobacter calcoaceticus RAG-1 on hexadecane. J Bacteriol 148: 51–57. Stach JEM, Burns RG (2002) Enrichment versus biofilm culture: A functional and phylogenetic comparison of polycyclic aromatic hydrocarbon-degrading microbial communities. Environ Microbiol 4: 169–182.

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Stewart PS, Franklin MJ (2008) Physiological heterogeneity in biofilms. Nat Rev Microbiol 6: 199–210. Webb JS, Givskov M, Kjelleberg S (2003) Bacterial biofilms: Prokaryotic adventures in multicellularity. Curr Opin Microbiol 6: 578–585. Whyte LG, Slagman SJ, Pietrantonio F, Bourbonnie`re L, Koval SF, Lawrence JR, Inniss WE, Greer CW (1999) Physiological adaptations involved in alkane assimilation at a low temperature by Rhodococcus sp. Strain Q15. Appl Environ Microbiol 65: 2961–2968. Wick LY, De Munain AR, Springael D, Harms H (2002) Responses of Mycobacterium sp. LB501T to the low bioavailability of solid anthracene. Appl Microbiol Biotechnol 58: 378–385. Wick LY, Pasche N, Bernasconi SM, Pelz O, Harms H (2003) Characterization of multiple-substrate utilization by anthracene-degrading Mycobacterium frederiksbergense LB501T. Appl Environ Microbiol 69: 6133–6142. Willison JC (2004) Isolation and characterization of a novel sphingomonad capable of growth with chrysene as sole carbon and energy source. FEMS Microbiol Lett 241: 143–150. Wolfaardt GM, Lawrence JR, Robarts RD, Caldwell DE (1995) Bioaccumulation of the herbicide diclofop in extracellular polymers and its utilization by a biofilm community during starvation. Appl Environ Microbiol 61: 152–158. Zilber Kirschner I, Rosenberg E, Gutnick D (1980) Incorporation of 32P and growth of pseudomonad UP-2 on n-tetracosane. Appl Environ Microbiol 40: 1086–1093.

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47 Production and Roles of Biosurfactants and Bioemulsifiers in Accessing Hydrophobic Substrates A. Perfumo . T. J. P. Smyth . R. Marchant . I. M. Banat* School of Biomedical Sciences, University of Ulster, Coleraine, County Londonderry, Northern Ireland, UK *[email protected] 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1502 2 Distribution of Biosurfactant-Producing Microorganisms in the Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1503 3 Rhamnolipid Biosurfactants in Pseudomonas aeruginosa . . . . . . . . . . . . . . . . . . . . . . . . . 1503 4 Surface-Active Lipids of Rhodococcus and Mycolate-Containing Microorganisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1506 5 Glycolipids of Alcanivorax in Marine Environments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1507 6 The Lipopeptide Biosurfactants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1508 7 High Molecular Mass Bioemulsifiers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1509 8 Research Needs and Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1510

K. N. Timmis (ed.), Handbook of Hydrocarbon and Lipid Microbiology, DOI 10.1007/978-3-540-77587-4_103, # Springer-Verlag Berlin Heidelberg, 2010

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Production and Roles of Biosurfactants and Bioemulsifiers

Abstract: Biosurfactants are one of the numerous adaptations of microorganisms metabolizing hydrocarbons and broadly represent a physiological response to specific requirements encountered by the cells depending on their environment. Some bacteria have developed the ‘‘pseudosolubilization’’ strategy to gain access to poorly soluble substrates and therefore produce highly dynamic low-molecular mass biosurfactants characterized by the capability to self-assembly in micelles, hemi-micelles or aggregates. Other bacteria interact with hydrocarbons directly by means of wall-bound biosurfactants that confer on the cell surface the appropriate hydrophobicity. High molecular mass bioemulsifiers in general adsorb tightly and cover the hydrocarbons thus dramatically increasing their apparent solubility. Although a variety of mechanisms and specializations distinguish biosurfactants, they all share a few essential traits. They are part of the process of interaction of microbial cells with surfaces as well as substrates. In addition to working on hydrocarbon solubilization they also act at the level of the cellular outer membrane allowing temporary and reversible modifications that control substrate access by the cells. As a consequence, a ‘‘substrate effect’’ can often be observed. Although the biosynthetic mechanisms for most remain unclear, substrates seem able to influence structural variations that make the biosurfactants particularly active towards the same substrate. Finally, many biosurfactants and their producing strains stimulate the growth of differently specialized bacteria which suggests that these molecules play a vital role in the interaction between the oil-degrading microbial communities and their environment.

1

Introduction

A large number of hydrophobic compounds continuously enter the environment, either as natural products originating from animals, plants and microorganisms such as steroids, terpenes and waxes or pollutants produced by anthropogenic activities such as hydrocarbons, petroleum and its derivatives. These substrates are degraded by microorganisms which have the ability to colonize almost all ecological niches as a result of their metabolic versatility and adaptability to different carbon sources. Many of these substances are characterized by low aqueous solubility and high solid-water distribution ratios (Johnsen et al., 2005). This limits their interaction with microbial cells which principally use molecules that are dissolved in the water phase. Substrate bioavailability is therefore an important factor, affected by complex processes in which many factors, not only physical and chemical substrate characteristics, play a significant role. They include for example the environment or matrix (e.g., water, soil, sediment, organic matter, etc.), the kinetic parameters (e.g., diffusion and flow rate, mass transfer, spatial separation between cells and substrate, etc.) as well as the intrinsic physiological properties of the cells (Johnsen et al., 2005). Microorganisms have adopted different strategies to enhance the bioavailability and gain access to hydrophobic compounds, such as hydrocarbons, including (1) biosurfactantmediated solubilization, (2) direct access of oil drops and (3) biofilm-mediated access (Hommel, 1990). The production of biosurfactants and bioemulsifiers is generally involved, although to different degrees, in all the above strategies. Biosurfactant structural uniqueness resides in the coexistence of a hydrophilic (a sugar or peptide) and a hydrophobic (fatty acid chain) domain in the same molecule, which allows them to occupy the interface of mixed phase systems (e.g., oil/water, air/water, oil/solid/water) and consequently to alter the forces governing the equilibrium conditions. This is the

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prerequisite for a broad range of surface activities to take place including emulsification, dispersion, dissolution, solubilization, wetting and foaming (Banat et al., 2000; Desai and Banat, 1997 (See > Chapter 27 and > 28, Vol. 5, Part 2)). Moreover biosurfactants seem to confer an essential evolutionary advantage allowing microorganisms to grow under specific conditions as evidenced by their wide distribution across the eubacterial and archeal domains (Bodour et al., 2003). This chapter will discuss how biosurfactants facilitate the hydrocarbon utilization in some representative hydrocarbon degrading-bacteria with emphasis on the diversity of these molecules (e.g., structural variants, physical and chemical properties and biosynthesis conditions) as specific response to needs, life-style and environmental niche of different microorganisms.

2

Distribution of Biosurfactant-Producing Microorganisms in the Environment

Biosurfactant-producing microorganisms are ubiquitous, inhabiting both water (sea, fresh water, groundwater) and land (soil, sediment, sludge) as well as extreme environments (e.g., oil reservoirs), and thriving at a wide range of temperatures, pH values and salinity. They can also be isolated from undisturbed environments where they have physiological roles not involving the solubilization of hydrophobic pollutants e.g., antimicrobial activity, biofilm formation or processes of motility and colonization of surfaces (Van Hamme et al., 2006). However, it is among the hydrocarbon-degrading microbial communities that the capability to produce biosurfactants is most widespread. Hydrocarbon-degrading bacterial populations are generally dominated by a few main bacterial genera such as Pseudomonas, Bacillus, Sphingomonas and Actinobacteria in soils and sediments, and Pseudoalteromonas, Halomonas, Alcanivorax and Acinetobacter in marine ecosystems. It is not surprising therefore that a lot of biosurfactant or bioemulsifier-producers belong to these same genera. An estimate of the frequency of biosurfactant-producing strains within a microbial population cannot be easily determined as it depends on the experimental procedures used. It has been reported that 2–3% of screened populations in uncontaminated soils are biosurfactant-producing microorganisms which increases to 25% in polluted soils (Bodour et al., 2003). On the other hand, enrichment culture techniques specific for hydrocarbon-degrading bacteria may lead to a much higher detection of biosurfactant-producers with estimates up to 80% (Rahman et al., 2002). However, the small number of investigations and a still naı¨ve study methodology limit our current understanding of biosurfactant-producing microorganisms so that occurrence and distribution in the environment are likely underestimated.

3

Rhamnolipid Biosurfactants in Pseudomonas aeruginosa

Rhamnolipids are biosurfactants exclusive to Pseudomonas aeruginosa although they have been recently reported to be produced by the phylogenetically related species Pseudomonas chlororaphis (Gunther et al., 2005). They are in the glycolipid family and their distinctive trait is the presence of one or two rhamnose sugar units linked to one or two units of b-hydroxy-fatty acids with typical chain length ranging from C8 to C12. Rhamnolipids are usually produced as a mixture of numerous congeners with the di-rhamnolipid-C10-C10 being the predominant form (> Fig. 1a,b). They are secondary metabolites whose production occurs constitutively at

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Production and Roles of Biosurfactants and Bioemulsifiers

. Figure 1 Structures of selected biosurfactants and bioemulsifiers from the most representative hydrocarbon-degrading microorganisms. L-rhamnosyl-3-hydroxydecanoyl-3-hydroxydecanoate (mono-rhamnolipid-C10-C10) (a) and L-rhamnosyl-L-rhamnosyl-3-hydroxydecanoyl-3hydroxydecanoate (di-rhamnolipid-C10-C10) (b) from Pseudomonas aeruginosa; trehalosedicorynomycolate (c) and trehalose-tetraester (d) from Rhodococcus sp.; glucolipid from Alcanivorax borkumensis (e); surfactin from Bacillus sp. (f); emulsan from Acinetobacter RAG-1 (g).

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low levels on both water-soluble and hydrophobic substrates with the concentration increasing during the late exponential-early stationary phase as a consequence of the higher cell density. The expression of the rhlAB operon and rhlC gene encoding the two rhamnosyltransferases responsible for the biosynthesis is under the control of the rhlR/rhlI genes, whose regulation is integrated into the quorum-sensing system (Soberon-Chavez et al., 2005). Most of our knowledge of rhamnolipid biosurfactants derives from investigations carried out in shaken-liquid cultures supplemented with hydrophobic compounds. In such conditions rhamnolipids are released into the medium as monomers that aggregate into micelles and larger vesicles when the concentration reaches and exceeds a critical level known as the critical micelle concentration (CMC). Hydrocarbons become incorporated within the hydrophobic core of micelles and this effectively enhances their dispersion into the aqueous phase and hence their bioavailability for uptake by cells. This process has been largely studied with alkanes as model substrates and is referred to as ‘‘micelle solubilization’’ or ‘‘pseudosolubilization’’ (> Figs. 2 and > 3) (Zhang and Miller, 1992). In addition rhamnolipids act on the cell outer membrane modifying the hydrophilic nature by inducing the release of lipopolysaccharide components with a subsequent increase of surface hydrophobicity. This enhances the affinity of cells for the hydrophobic substrate facilitating access to the finely dispersed oil droplets (Al-Tahhan et al., 2000). This same mode of action also seems to be involved in the uptake of solid and non-aqueous phase liquid (NAPL) polycyclic aromatic hydrocarbons. Small amounts of surface-exposed biosurfactants and polymeric substances available when cell numbers are low allow initial growth at the interface between water and oil and is essential to start the mass transfer of the substrate into the bulk aqueous phase and provide an initial carbon input that further supports the development of a bigger competent bacterial community (Garcı´a-Junco et al., 2001). In solid phases such as soil, sediment, sand or organic matter, where the hydrophobic substrates can be present as solid particles or liquid film adsorbed onto the matrix or sequestered in pores and the bacteria are heterogeneously distributed, the production of biosurfactants first increases the substrate accessibility and reduces its spatial separation from the degrading cells. Rhamnolipids are believed to aggregate at the solid interface forming single mono-layers known as hemi-micelles capable of desorbing the hydrophobic substrates thus increasing their bioavailability in the aqueous phase (Volkering et al., 1998). However,

. Figure 2 Mechanism of hydrocarbon solubilization within biosurfactant micelles. At low concentration, biosurfactants occur as monomers at the interface between the aqueous and hydrocarbon phases (a). When the concentration increases and the space available decreases, biosurfactants tend to arrange into aggregates (b) up to a point called the ‘‘critical micelle concentration’’ at which micelles are formed trapping the hydrocarbons into their hydrophobic core (c). Once dispersed, hydrocarbons become more available to uptake by the cells.

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. Figure 3 Photomicrograph of a mixed population of bacteria growing in the presence of hydrocarbons. Cells tend to occur at the interface between the aqueous and hydrophobic phases and a micelle (see arrow) can be also observed.

experimental conditions may present a quite different scenario. In soil microcosms where hydrocarbons are well mixed and the bacteria homogenously distributed over the available surface, the distance between substrate and cells is artificially reduced making direct contact the preferred uptake mode. Although the expression of rhamnolipids is constitutive, their contribution to hydrocarbon degradation is probably marginal (Holden et al., 2002).

4

Surface-Active Lipids of Rhodococcus and Mycolate-Containing Microorganisms

Rhodococcus is one of the most active hydrocarbon-degrading bacterial genera due to a broad set of potent enzymes, oxygenases in particular, and robust cellular physiology. Their ability to produce biosurfactants is one of the numerous adaptations induced by the presence of hydrocarbons. Uptake of alkanes occurs via direct access to large oil drops by the cells with biosurfactants mediating the interaction. This implies that biosurfactants are synthesized in a growthassociated manner and that most of them remain wall-bound conferring the cell surface with that essential hydrophobicity through which the contact with the substrate can take place. Other components of the cell wall such as glycolipids, mycolic and fatty acids also contribute to the remarkably hydrophobic nature of Rhodococcus and the CMN group (Corynebacterium, Mycobacterium and Nocardia). An additional effect is that wetting of the cell surface is hampered and this further supports the access to hydrocarbons. Only a small portion of biosurfactants, estimated to be approximately 10%, are in fact released into the culture medium to help the preliminary emulsification of the oil (Lang and Philp, 1998).

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Growth and metabolic activities of Rhodococcus therefore occur at the